Slippery liquid-infused porous surfaces that prevent microbial surface fouling

ABSTRACT

The present invention provides polymer-based slippery liquid-infused porous surfaces (SLIPS) that can prevent adhesion and colonization by fungal and bacterial pathogens and also kill and/or attenuate the colonization and virulence of non-adherent pathogens in surrounding media. The present approach exploits the polymer and liquid oil phases in these slippery materials to sustain the release of small molecules such as a broad-spectrum antimicrobial agent, an antifungal agent, an antibacterial agent, an agent that modulates bacterial or fungal quorum sensing, an agent that attenuates virulence, or a combination thereof. This controlled release approach improves the inherent anti-fouling properties of SLIPS, has the potential to be general in scope, and expands the potential utility of slippery, non-fouling surfaces in both fundamental and applied contexts.

CROSS-REFERENCE TO RELATED APPLICATIONS

This application claims priority from U.S. patent application Ser. No.15/471,628, filed Mar. 28, 2017, and U.S. Provisional Patent ApplicationNo. 62/314,282, filed Mar. 28, 2016. Both are incorporated by referenceherein to the extent that there is no inconsistency with the presentdisclosure.

STATEMENT REGARDING FEDERALLY SPONSORED RESEARCH OR DEVELOPMENT

This invention was made with government support under N00014-07-1-0255and N00014-14-1-0791 awarded by the US Navy/ONR; DMR1121288 DMR0832760awarded by the National Science Foundation; and AI092225 awarded by theNational Institutes of Health. The government has certain rights in theinvention.

BACKGROUND OF THE INVENTION

Synthetic surfaces that are resistant to fouling by aqueous media,organic fluids, or biological organisms are critical in a broad range ofindustrial, commercial, and biomedical contexts. Surfaces that aresuperhydrophobic, superoleophobic, or superomniphobic, for example, forma basis for the design of self-cleaning and antifogging materials,anti-corrosive interfaces, and stain-resistant textiles, and haveenabled new strategies for the transport and manipulation of complexfluids, including approaches to oil recovery and oil/water separation(see Liu et al., Chem. Soc. Rev. 2010, 39, 3240; Banerjee et al., Adv.Mater. 2011, 23, 690; Yao et al., Adv. Mater. 2011, 23, 719; Liu et al.,Ann. Rev. Mater. Res. 2012, 42, 231; Campoccia et al., Biomaterials2013, 34, 8533; Ueda et al., Adv. Mater. 2013, 25, 1234; Bellanger etal., Chem. Rev. 2014, 114, 2694; Genzer et al., Science 2000, 290, 2130;Tuteja et al., Science 2007, 318, 1618; Chu et al., Chem. Soc. Rev.2014, 43, 2784; and Deng et al., Science 2012, 335, 67).

Slippery liquid-infused porous surfaces (SLIPS) are an emerging class ofsynthetic materials that exhibit unique and robust antifouling behavior.These materials are fabricated by infusion of viscous oils into poroussurfaces, yielding interfaces that allow other fluids to slide off withsliding angles sometimes as low as 2°. This slippery behavior arisesfrom an ability to host and maintain thin films of oil at theirsurfaces, placing a premium on chemical compatibility between the matrixand the oil and revealing design criteria that can be exploited tomanipulate the behaviors of contacting fluids (e.g., to tune slidingangles and velocities or create responsive surfaces that allow controlover these and other interfacial behaviors). Surfaces and coatings thatexhibit these characteristics have enabled the design of new anti-icingsurfaces, slippery containers for the dispensing of commercial liquidsand gels, and new liquid-infused interfaces that are resistant tobiofouling in complex aqueous, biological, and marine environments.

Recent reports on alternative approaches to the development of SLIPShave enabled the design of new classes of synthetic and highly‘slippery’ anti-fouling materials that address practical limitationsexhibited by conventional non-wetting (e.g., superhydrophobic) surfaces,and introduce new principles for the design of robust, injury-tolerant,and mechanically compliant synthetic anti-fouling surfaces (see Wong etal., Nature 2011, 477, 443; Grinthal et al., Chem. Mater. 2014, 26, 698;Epstein et al., Proc. Natl. Acad. Sci. U.S.A. 2012, 109, 13182; Yao etal., Nat. Mater. 2013, 12, 529; Liu et al., Adv. Mater. 2013, 25, 4477;Smith et al., Soft Matter 2013, 9, 1772; Vogel et al., Nat. Commun.2013, 4; Huang et al., ACS Macro Lett. 2013, 2, 826; Leslie et al., Nat.Biotechnol. 2014, 32, 1134; Glavan et al., Adv. Funct. Mater. 2014, 24,60; Wei et al., Adv. Mater. 2014, 26, 7358; Yao et al., Adv. Mater.2014, 26, 1895; and Zhang et al., Adv. Funct. Mater. 2014, 24, 1074.)

Reports by Aizenberg, Levkin, and others demonstrate that SLIPS can bedesigned to resist fouling by bacteria and other marine organisms thatcan colonize and form biofilms on biomedical devices or commercial andindustrial equipment (see Epstein et al., Proc. Natl. Acad. Sci. U.S.A.2012, 109, 13182; Leslie et al., Nat. Biotechnol. 2014, 32, 1134; Howellet al., ACS Appl. Mater. Inter. 2014, 6, 13299; Li et al., ACS Appl.Mater. Inter. 2013, 5, 6704; and Xiao et al., ACS Appl. Mater. Inter.2013, 5, 10074). Those studies suggest that appropriately designedliquid-infused surfaces can resist the attachment, colonization, andorganization of communities of these organisms in ways that exceed thoseexhibited by some conventional anti-fouling surfaces (such as surfacesmodified with polyethylene glycol and non-wetting superhydrophobicsurfaces, etc.), even in complex media with proteins, surfactants, or athigh ionic strengths typical of environmental conditions encountered inmany applied and biologically relevant contexts.

While these past studies represent outstanding progress toward thedesign of new anti-fouling materials with superior functionalproperties, existing SLIPS do not completely inhibit colonization, andfundamental questions remain regarding both the long-term stabilities ofthese materials and the ability of bacteria or other microorganisms toadapt to or breach the infused liquid barriers that confer slipperycharacter to establish ‘beachheads’ on underlying surfaces that couldenable colonization and bio-fouling. In addition, and of particularinterest in the context of potential biomedical applications of thesenew materials, SLIPS can currently do little to influence the behaviorsof planktonic microorganisms—that is, while SLIPS can substantiallyprevent the adhesion of pathogenic microorganisms or the formation ofmicrobial biofilms on treated surfaces, they cannot prevent the growthor proliferation of those organisms in solution, prevent them fromcolonizing other nearby surfaces, or stop them from engaging in otherbehaviors (e.g., toxin production) that could lead to infection andother associated burdens.

The present invention addresses these issues and allows for thefunctional properties and potential applications of SLIPS to besignificantly expanded by adopting design strategies that leverage thepotential of the porous matrices and the infused and lubricating oils inthese materials as depots for the storage and subsequent release ofbioactive agents. In particular, small-molecule anti-microbial agentscan be stored in the porous matrices or dissolved and stored in afugitive oil phase without compromising the ‘slippery’ characteristic ofthe material, thereby providing new approaches to the design ofmulti-functional or dual-action SLIPS with improved antimicrobialproperties. Provided that the embedded agent can also diffuse into theoil phase and/or from the oil phase into surrounding aqueous media, thisapproach also offers opportunities to design anti-fouling SLIPS thatcould kill or influence the behaviors of planktonic microorganisms. In abroader and more general context, the ability to store and control therelease of small molecules or other agents from SLIPS allows for a widerange of other new applications for these liquid-infused materials.

SUMMARY OF THE INVENTION

Conventional oil-infused ‘slippery’ surfaces can prevent surface foulingowing to their slippery character, which reduces the ability of bacteriaor other substances from adhering to the surface, but cannot otherwiseinfluence events that occur in that surrounding media or change theproperties of that surrounding media. The present invention relates tomethods for the design of multifunctional slippery surfaces that enablethese surfaces to exert influences and affect new and desired behaviorsin surrounding environments. The present invention is generally directedtoward the design of antifungal and antibacterial polymer-based SLIPSthat inhibit microbial adhesion and/or promote the sustained release ofsmall-molecule compounds, such as a broad-spectrum antimicrobial agent,an antifungal agent, an antibacterial agent, an agent that modulatesbacterial or fungal quorum sensing, an agent that attenuates virulence,or combinations thereof.

In an embodiment, the present invention provides polymer-based slipperyliquid-infused porous surfaces (SLIPS) that can prevent adhesion andcolonization by fungal and bacterial pathogens and also kill and/orattenuate the colonization and virulence of non-adherent pathogens insurrounding media. The present approach exploits the polymer and liquidoil phases in these slippery materials to sustain the release ofsmall-molecule compounds, such as antimicrobial agents, antifungalagents, virulence attenuating agents, and bacterial or fungal quorumsensing agents. This controlled release approach improves the inherentanti-fouling properties of SLIPS, has the potential to be general inscope, and expands the potential utility of slippery, non-foulingsurfaces in both fundamental and applied contexts.

One embodiment of the present invention provides a slipperyliquid-infused porous surface (SLIPS) that controllably releases amolecule, wherein the slippery oil-infused surface comprises: a) aporous matrix having nanoscale or microscale porosity; b) an oilcovering at least a portion of the porous matrix, wherein the oil atleast partially fills the pores of the porous matrix; and c) one or moresmall-molecule compounds able to reduce, inhibit, or modulate thebehaviors of non-adherent pathogens in surrounding media, wherein theone or more small-molecule compounds are located on the surface orwithin said porous matrix, within said oil, or both, and wherein theslippery oil-infused surface is able to controllably release the one ormore small-molecule compounds when the slippery oil-infused surface isimmersed into said media. In an embodiment, the porous matrix hasnanoscale porosity.

As used herein, “small-molecule compounds” refer to compounds having amolecular weight of approximately 900 daltons or less, preferablyapproximately 700 daltons or less, preferably approximately 500 daltonsor less, or preferably approximately 300 daltons or less. It isunderstood that the chemical structure of the molecule will influenceits solubility in the oil phase, its solubility in the water phase, andits interactions with the polymer matrix in ways that will influence,and which can be used to modulate, its release profile into thesurrounding media. In an embodiment, the small-molecule compound issoluble to very soluble in water (at least 3.3 g/100 g H₂O). In anembodiment, the small-molecule compound is sparingly soluble in water(0.1 to 3.3 g/100 g H₂O). In an embodiment, the small-molecule compoundis slightly soluble in water (0.01 to 0.1 g/100 g H₂O). In anembodiment, the small-molecule compound is practically insoluble (lessthan 0.01 g/100 g H₂O). In an embodiment, the small-molecule compound isnot a polypeptide. In an embodiment, the small-molecule compounds havedrug-like characteristics such as good absorption, distribution,metabolism, excretion and toxicity (ADMET) profiles as known in the art(see, for example, Lipinski, Journal of Pharmacological andToxicological Methods 2000, 44: 235-249).

As used throughout this invention, preferably the one or moresmall-molecule compounds are able to reduce, inhibit, or modulate thebehaviors of non-adherent pathogens in the surrounding media. Asnon-limiting examples, the one or more small-molecule compounds can killor otherwise reduce at least a number of the pathogens, slowreproduction or growth of least a portion of pathogens, or modulatebehavior such as preventing or reducing the ability of pathogens tocommunicate with each other. In an embodiment, the one or moresmall-molecule compounds comprise an antimicrobial agent, an antifungalagent, an antibacterial agent, an agent that modulates bacterial orfungal quorum sensing, an agent that attenuates virulence, or acombination thereof. In an embodiment, the one or more small-moleculecompounds is a natural or synthetic antibiotic agent, natural orsynthetic antifungal agent, quorum sensing modulator, or a combinationthereof. In an embodiment, the one or more small-molecule compoundscomprise one or more anti-microbial peptides having a molecular weightof 900 daltons or less. In another embodiment, the one or moresmall-molecule compounds do not include any peptides. Optionally, theone or more small-molecule compounds is hydrophobic.

Preferably, the one or more small-molecule compounds are able to reduce,inhibit, or modulate fungal and bacterial pathogens including, but notlimited to, Candida species, Aspergillus species, Cryptococcus species,Histoplasma species, Helicobacter species, Neisseria species,Pneumocystis species, Stachybotrys species, Pseudomonas species,Escherichia species, Streptococcus species and Staphylococcus species.

In further embodiments, the one or more small-molecule compounds isselected from the group consisting of acyl L-homoserine lactone (AHL)derivatives, aminobenzimidazole (ABI) derivatives, and combinationsthereof. Classes of useful small-molecule drugs are modulators andparticularly antagonists of bacterial quorum sensing. A number of suchsmall-molecule modulators are known in the art and several exemplaryquorum sensing modulators are illustrated below. Eibergen et al.,ChemBioChem 2015, 16:2348-2356, reports among others certain classes ofquorum sensing antagonists designated PHL's and POHL's therein asexemplified by compounds A and B shown below. Moore et al., J. Amer.Chem. Soc. 2015, 137:14626-14639 reports among others AHL mimics whichare quorum sensing antagonists such as compound C and certain non-AHLmodulators such as compound D (shown below). O'Reilly et al., ACSInfect. Dis. 2016, 2:32-38, for example, reports among othershydrolytically stable LasR antagonists such as compounds E and F (shownbelow). Starkey et al., PLoS Pathog. 2014, 10, e100432,1 reportcompounds which disrupt quorum sensing such as compound G (shown below).Frei et al., Angewandte Chemie 2012, 124:5316-5319 report2-aminobenzimidazoles, such as compound H (shown below), which inhibitand disperse biofilms. Each of these references is incorporated byreference herein in its entirety for descriptions of quorum sensingmodulators, particularly antagonists of quorum sensing, includingdescriptions of their preparation and their activities. U.S. Pat. Nos.8,815,943; 8,624,063; 8,367,680; 8,269,024; 7,910,622; and 7,642,285relate to small molecule quorum sensing modulators useful in the methodsof the present invention.

In further embodiments, the one or more small-molecule compounds isselected from the group consisting of:

or combinations thereof.

One embodiment of the invention is based on SLIPS fabricated by theinfusion of a hydrophobic liquid oil into nanoporous polymer multilayersfabricated by reactive/covalent layer-by-layer assembly, such asdescribed in Manna et al., Adv. Mater. 2015, 27, 3007; Buck et al., Adv.Mater. 2007, 19, 3951; Buck et al., Polym. Chem. 2012, 3, 66; and Mannaet al., Adv. Funct. Mater. 2015, 25, 1672. The present invention furtherdemonstrates that (i) these polymer-based SLIPS can substantiallyprevent surface fouling, including biofilm formation, by several typesof common fungal and bacterial human pathogens, and (ii) that biofilmformation on the SLIPS-coated surfaces of planar objects andpolymer-based catheter tubes can be reduced further by using porouspolymer matrices loaded with one or more antifungal or antibacterialagents, such as triclosan, a model broad-spectrum antimicrobial agent.

The present invention also demonstrates that the release of anantimicrobial agent into the surrounding media can be used toefficiently and effectively kill planktonic microorganisms present insurrounding media and thereby prevent biofilm formation on neighboringuncoated surfaces. Without being bound by theory, it is believed theSLIPS of the present invention sustain the release of the one or moresmall-molecule compounds by the partitioning of the one or moresmall-molecule compounds from the porous polymer matrix into the liquidoil phase and from the liquid oil phase into the surrounding aqueousmedia. The experimental results presented introduce new principles thatcould prove useful for the design of multi-functional slippery surfaceswith improved anti-fouling properties. However, it should be noted thatthe one or more small-molecule compounds can be loaded into the oilitself, and thus the controlled release is determined by thepartitioning of the small-molecule compound from the oil into thesurrounding aqueous media. Alternatively, the small-molecule compoundsmay travel along pathways or channels formed at the interphase of theoil infused porous matrix and other regions.

In an embodiment, the surrounding media is an aqueous media where thesurface may encounter fungi, bacteria, and/or other microorganisms.Types of surrounding media include, but are not limited to, salt waterenvironments (such as sea water or saline solutions), fresh waterenvironments (such as swamp water or fresh lake water), andphysiological or physiologically relevant media (including but notlimited to phosphate-buffered saline solutions, TRIS-buffered salinesolutions, HEPES-buffered saline solutions, Ringer's solution, cellculture media as known in the art, blood or blood plasma, and otherbodily fluids). Preferably, the surrounding media does not promote theleaching of the oil or degrade the oil phase of the SLIPS, or does so ata slow rate.

In an embodiment, the present invention provides a multilayer filmcomprising one or more bilayers infused with an oil, wherein eachbilayer comprises an optionally functionalized first polymer layer incontact with a second polymer layer, and wherein the multilayer film hasa nanoscale or microscale porosity. Preferably, the multilayer film hasnanoscale porosity. The infusion of the oil into at least a portion ofthe rough or porous surfaces of the multilayer film causes other liquidsplaced in contact with the multilayer film to slide off the multilayerfilm or a surface coated with the multilayer film. Additionally, themultilayer film comprises one or more small-molecule compounds able tobe controllably released from the multilayer film into the surroundingmedia.

In an embodiment, the present invention provides a method forfabricating a slippery liquid-infused porous surface (SLIPS) able toreduce or inhibit non-adherent pathogens in surrounding media, themethod comprising the steps of: a) forming a porous matrix on asubstrate, wherein said porous matrix has nanoscale or microscaleporosity; b) exposing the porous matrix to an oil, wherein said oilcoats at least a portion of the porous matrix and said oil at leastpartially fills the pores of at least a portion of said porous matrix;and c) loading one or more small-molecule compounds onto or into saidporous matrix or into said oil. Optionally, the one or moresmall-molecule compounds is loaded onto or into the porous matrix, orwithin the oil itself, prior to exposing the porous matrix to the oil.Alternatively, the one or more small-molecule compounds are loadedwithin the oil after exposing the porous matrix to the oil. In a furtherembodiment, an additional one or more small-molecule compound is loadedinto the oil when levels of small-molecule compounds drop below adesired level, such as from prolonged use of the SLIPS. The newly addedone or more small-molecule compounds can be the same or different thanthe original small-molecule compounds. Preferably, the one or moresmall-molecule compounds is able reduce, inhibit, or modulate thebehaviors of said pathogens upon contact with said pathogens.

In another embodiment, the present invention provides a method forfabricating a slippery liquid-infused porous surface (SLIPS) multilayerfilm able to reduce or inhibit non-adherent pathogens in surroundingmedia, where the multilayer film comprises one or more bilayers, themethod comprising the steps of: a) exposing a surface of a substrate toa first solution comprising a first polymer wherein a layer of the firstpolymer is deposited on at least a portion of the substrate; b) exposingthe substrate to a second solution comprising a second polymer whereinthe second polymer reacts with the first polymer layer and a layer ofthe second polymer is deposited on at least a portion of the firstpolymer layer, thereby forming a bilayer in the multilayer film; c)exposing the substrate to an oil wherein said oil coats at least aportion of the multilayer film and said oil at least partially fills thepores of at least a portion of said multilayer film; and d) loading oneor more small-molecule compounds onto or into said one or more bilayersor into said oil. Preferably, the one or more small-molecule compoundsis able to reduce, inhibit, or modulate the behavior of said pathogensupon contact with the pathogens.

Optionally, the one or more small-molecule compounds are loaded onto orinto the one or more bilayers, or within the oil itself, prior toexposing the substrate to the oil. Alternatively, the one or moresmall-molecule compounds are loaded within the oil after exposing thesubstrate to the oil. When administered to the one or more bilayers, theone or more small-molecule compounds can be deposited on, or within, theoutermost bilayer. Alternatively, the one or more small-moleculecompounds can be deposited on, or within, multiple bilayers or even allof the bilayers. In a further embodiment, an additional one or moresmall-molecule compound is loaded into the oil when levels ofsmall-molecule compounds drop below a desired level, such as fromprolonged use of the multilayer film. The newly added one or moresmall-molecule compounds can be the same or different than the originalsmall-molecule compounds.

The fabrication method relating to the multilayer film optionallycomprises a rinsing step comprising exposing or washing the substratewith a rinse solvent or solution each time step a) is performed and eachtime step b) is performed. In an embodiment, a fresh rinse solvent orsolution is employed for each rinsing step. In a further embodiment, thesame rinse solution is re-used for each rinsing step.

Preferably, steps a) and b) relating to the multilayer film are repeatedone or more times until the multilayer film reaches the desiredthickness or desired number of layers before the substrate is exposed tothe oil, where each cycle deposits a new bilayer on the substrate. Inspecific embodiments, the multilayer polymer film comprises more thanone bilayer. In a further embodiment, steps a) and b) are repeated 2 ormore times, 5 or more times, 10 or more times, 20 or more times, 30 ormore times, 50 or more times, or 100 or more times. The substrate can beexposed to the solutions containing the polymer solutions using methodsknown in the art, including but not limited to, dip coating.

The substrate can be any material able to support the formation of thenanoporous or microporous porous matrix, including but not limited toglass, metals and plastics. The substrate can include curved andirregularly shaped three dimensional surfaces, as well as completelysolid surfaces and mesh surfaces (e.g., having a porosity between 100 μmand 250 μm). For example, the substrate can be the interior of a tube orcontainer for a liquid or gel where it is undesirable for the contentsof the tube or container to stick or adhere to the surface. The porousmatrix, first polymer layer, second polymer layer, and oil are thereforeselected so that the liquid or gel has reduced adhesion to thecontainer. Alternatively, the substrate can be a display of a sensorwhere the degree or extent to which a liquid adheres to the substrateindicates the presence of a substance in the liquid.

A further embodiment of the invention provides for patterning thesubstrate so that the multilayer film is formed on a first specifiedportion of the substrate, thereby creating a substrate having one ormore “slippery” regions and one or more “sticky” regions. A portion ofthe multilayer film on the first specified portion of the substrate isfurther functionalized with an amine or hydroxyl group having theformula R—NH₂ or R—OH, where R is hydrophobic. In a further embodiment,a second specified portion of the substrate is not covered by the oilinfused porous matrix, or, alternatively, a portion of the one or morebilayers on the second specified portion of the substrate is furtherfunctionalized with an amine or hydroxyl group having the formula R—NH₂or R—OH, where R is hydrophilic.

Additionally, in a further embodiment, a portion of the one or morebilayers on the first specified portion of the substrate is furtherfunctionalized with an amine or hydroxyl group having the formula R—NH₂or R—OH, where R is hydrophobic, a second specified portion of thesubstrate is not covered by the oil infused multilayer film, and a thirdportion of the substrate is covered by a bilayer where a portion of theone or more bilayers on the third specified portion of the substrate isfurther functionalized with an amine or hydroxyl group having theformula R—NH₂ or R—OH, where R is hydrophilic.

The first and second polymer layers of the bilayer can comprise anypolymers or combination of polymers able to form a stable bilayer andwhere the first polymer layer is optionally able to be functionalizedand the second polymer layer is optionally also able to befunctionalized (as described in U.S. Pat. No. 8,071,210). The chemicalreactivity of the functionalized bilayers provides means to tuneinteractions between the matrix and infused oil phases. Spatial controlover the functionalization can be used to create SLIPS with regionsdevoid of oil that can prevent or arrest the sliding of aqueous fluids,extract samples of liquid from contacting media, or provide control overthe trajectories of sliding droplets. Preferably, the first polymerlayer is covalently cross-linked with the second polymer layer. Infurther embodiments, the bilayers are reacted with small chemical groupscontaining a hydrophobic or hydrophilic amine to further functionalizethe bilayer (i.e., to install secondary surface functionality).

In an embodiment, the first polymer layer of the bilayer comprises afunctionalized azlactone having the formula:

wherein x is 0 or the integers 1 or 2; and each R¹ is independentlyselected from the group consisting of: hydrogen, alkyl groups, alkenylgroups, alkynyl groups, carbocyclic groups, heterocyclic groups, arylgroups, heteroaryl groups, alkoxy groups, aldehyde groups, ether groups,and ester groups, any of which may be substituted or unsubstituted. Inan embodiment, the first polymer layer comprises functionalizedpoly(vinyl-4,4-dimethylazlactone) (PVDMA). In an embodiment, the firstpolymer layer consists of functionalizedpoly(vinyl-4,4-dimethylazlactone) (PVDMA). In a further embodiment, thePVDMA is synthesized by free-radical polymerization of PVDMA withintentionally added cyclic azlactone-functionalized oligomer in anamount ranging from 1 wt % to 10 wt %, preferably between 5 wt % and 8wt %.

Useful functionalized azlactone polymers include, but are not limitedto, poly(vinyl-4,4-dimethylazlactone),poly(2-vinyl-4,4-dimethyl-2-oxazolin-5-one),poly(2-isopropenyl-4,4-dimethyl-2-oxazolin-5-one),poly(2-vinyl-4,4-diethyl-2-oxazolin-5-one),poly(2-vinyl-4-ethyl-4-methyl-2-oxazolin-5-one),poly(2-vinyl-4-dodecyl-4-methyl-2-oxazolin-5-one),poly(2-vinyl-4,4-pentamethy lene-2-oxazolin-5-one), poly(2-vinyl-4-methyl-4-phenyl-2-oxazolin-5-one),poly(2-isopropenyl-4-benzyl-4-methyl-2-oxazolin-5-one), orpoly(2-vinyl-4,4-dimethyl-1,3-oxazin-6-one). Useful azlactonefunctionalized polymers further include azlactone functionalizedpolyisoprenes and azlactone functionalized polybutadienes.

In an embodiment, the second polymer layer of the bilayer is optionallyfunctionalized and comprises an amine functionalized polymer, an alcoholfunctionalized polymer, or a thiol functionalized polymer. Creatingspecific functionalities with amine, alcohol, and thiol groups is aprocess well known in the art (for example, see Bioconjugate Techniques,2^(nd) Edition, 2008, Greg T. Hermanson). In embodiments, the secondpolymer layer comprises an optionally functionalized polymer selectedfrom the group consisting of poly(ethylene imine) (PEI), polylysine,pollyallylamine, poly(amidoamine) dendrimers, polyvinyl alcohol, polyhydroxyl ethyl methacrylate, poly(methacrlic acid) functionalized withcysteamine, and linear and hyperbranched and dendritic polymersfunctionalized with primary amines, hydroxyl groups, or thiol groups.

In embodiments, the second polymer layer comprises a polymer, which isoptionally functionalized, selected from the group consisting ofpolyolefins, poly(alkyls), poly(alkenyls), poly(ethers), poly(esters),poly(imides), polyamides, poly(aryls), poly(heterocycles), poly(ethyleneimines), poly(urethanes), poly(α,β-unsaturated carboxylic acids),poly(α,β-unsaturated carboxylic acid derivatives), poly(vinyl esters ofcarboxylic acids), poly(vinyl halides), poly(vinyl alkyl ethers),poly(N-vinyl compounds), poly(vinyl ketones), poly(vinyl aldehydes) andany combination thereof. In an embodiment, the second polymer layercomprises poly(ethylene imine) (PEI).

For some embodiments, it may be desirable to further functionalize aportion of the one or more bilayers. This can be achieved, for example,by reacting a portion of any residual functional groups in the one ormore bilayers with an amine group or hydroxyl group, or by reacting aportion of the first or second polymer with an amine reactive group orhydroxyl reactive group.

In an embodiment, at least a portion of the residual functional groupsin the bilayer is reacted with an amine or hydroxyl group having theformula R—NH₂ or R—OH, where R is hydrophobic or hydrophilic. Inembodiments, R is a substituted or unsubstituted C₁ to C₂₀ alkyl group,preferably a C₁ to C₁₂ alkyl group. In other embodiments, R is asubstituted or unsubstituted C₂ to C₂₀ alkenyl group, preferably a C₂ toC₁₂ alkenyl group. In further embodiments, at least a portion of theresidual functional groups in the bilayer is reacted with an amineselected from the group consisting of methylamine, ethylamine,propylamine, butylamine, pentylamine, hexylamine, heptylamine,octylamine, nonylamine, decylamine, and combinations thereof, preferablyn-propylamine, n-octylamine, or n-decylamine. In other embodiments, R isan alkyl group substituted with one or more hydroxyl groups or chargedgroups such as COO⁻ or NR₃ ⁺. In an embodiment, at least a portion ofthe residual functional groups in the bilayer is reacted with an aminosugar, amino alcohol, amino polyol, glucamine (preferably D-glucamine),dimethylaminopropylamine (DMAPA), and combinations thereof.

In an embodiment, the polymer of the first polymer layer is furtherfunctionalized with a hydrophobic (decylamine or propylamine) orhydrophilic (glucamine) primary amine-containing small molecule.

As used herein, “an oil” refers to any water-immiscible phase,preferably a non-polar, hydrophobic chemical substance which is a liquidat ambient temperature and which has no or very low solubility in water.Preferably, the oil is selected so as to either completely or partiallysolubilize the small-molecule compounds. The oil infused into the one ormore bilayers can be a synthetic oil or a natural oil, and is preferablya biocompatible oil. Preferably, the oil is selected from the groupconsisting of a silicone oil, a vegetable oil, a mineral oil, aperfluorinated oil, a thermotropic liquid crystal, an anisotropic oil,and combinations thereof. Suitable vegetable oils include, but are notlimited to, canola oil, coconut oil, olive oil, soybean oil andcombinations thereof. In some embodiments, silicone oil is selected dueto improved solubility with the one or more small-molecule compounds.

In an embodiment, a slippery liquid-infused porous surface (SLIPS)coating is provided comprising: a) a multilayer polymer film comprisingone or more bilayers where said multilayer polymer film has a nanoscaleor microscale porosity, wherein each bilayer comprises a first polymerlayer covalently linked with a second polymer layer; b) an oil selectedfrom the group consisting of a silicone oil, a vegetable oil, a mineraloil, a thermotropic liquid crystal, and combinations thereof, whereinsaid oil coats at least a portion of the multilayer polymer film andsaid oil at least partially fills the pores of at least a portion ofsaid multilayer polymer film; and c) one or more small-moleculecompounds able to reduce or inhibit non-adherent pathogens insurrounding media, wherein the one or more small-molecule compounds arelocated on the surface of the one or more bilayers, within the oil, orboth, wherein the multilayer film is able to controllably release aneffective amount of one or more small-molecule compounds when saidmultilayer film is immersed into said media. Preferably, the multilayerpolymer film of the coating has a thickness of 5 μm or less, andcomprises one or more PVDMA/PEI bilayers, which are furtherfunctionalized with a hydrophobic amine. Preferably, the multilayerpolymer film has nanoscale porosity.

A specific embodiment of the present invention provides a SLIPS designbased on the infusion of oils into nanoporous or microporous (preferablynanoporous) polymer coatings fabricated by reactive layer-by-layerassembly of polymer multilayers using branched poly(ethylene imine)(PEI) and the amine-reactive polymer poly(vinyl-4,4-dimethylazlactone)(PVDMA). In an embodiment, the multilayer film comprises one or morePVDMA/PEI bilayers, which are further functionalized with a decyl groupby reacting with n-decylamine and wherein the one or more bilayers areinfused with a silicone oil or an anisotropic thermotropic liquidcrystal.

One aspect of the invention provides thin multilayer polymer films andcoatings (e.g., equal to or less than 100 μm, equal to or less than 50μm, preferably less than or equal to 10 μm, preferably less than orequal to 5 μm). Preferably, the multilayer film comprises 2 or morebilayers, 5 or more bilayers, 10 or more bilayers, 20 or more bilayers,30 or more bilayers, 50 or more bilayers, or 100 or more bilayers.Preferably each first polymer layer alternates with the second polymerlayer. In embodiments, the multilayer films have a nanoscale ormicroscale porosity. Preferably, the multilayer films have nanoscaleporosity.

In an embodiment, SLIPS are infused with a thermotropic liquid crystal(an anisotropic oil) to generate sliding angles and velocities thatdepend critically upon the chemical compositions of contacting aqueousphases, revealing a novel ‘sliding’ basis for the sensing and naked-eyedetection of environmental analytes, including bacterial endotoxin(i.e., LPS) in aqueous media via visually apparent changes in dropletsliding speeds as a function of analyte concentration. Such LC-infusedSLIPS provide opportunities to design slippery surfaces that couldpermit active and external control over droplet adhesion and mobility.

In an embodiment, the present invention provides a method for reducing,inhibiting, or modulating the behaviors of non-adherent pathogens inmedia surrounding a substrate comprising the steps of: a) providing aslippery liquid-infused porous surface (SLIPS) on the substrate, saidslippery oil-infused surface comprising:

i) a porous matrix having nanoscale or microscale porosity;

ii) an oil covering at least a portion of the porous matrix, whereinsaid oil at least partially fills the pores of the porous matrix; and

iii) one or more small-molecule compounds able to reduce, inhibit, ormodulate the behaviors said pathogens upon contact with said pathogens,wherein the one or more small-molecule compounds are located on thesurface of said porous matrix, within said oil, or both; and

b) controllably releasing the one or more small-molecule compounds fromsaid slippery oil-infused surface into said media, wherein the one ormore small-molecule compounds contact the pathogens thereby reducing thenumber of pathogens, inhibiting the growth or colonization of thepathogens, or modulating the behaviors of the pathogens.

In a further embodiment, one or more additional small-molecule compoundsare loaded within the oil when the levels of small-molecule compoundsdrop below a desired level or when different small-molecule compoundsare desired. For example, a different anti-fungal or anti-bacterialcompound can be added to the SLIPS depending on which pathogens arecurrently present in the surrounding media. Alternatively, thesmall-molecule compounds can be replenished when a significant amount ofthe small-molecule compounds has been released into the surroundingmedia by the SLIPS during use. In an embodiment, the porous matrix is influid communication with a reservoir containing additional amounts ofthe oil, small-molecule compounds or both. When the amount of oil orsmall-molecule compounds at the surface of the SLIPS is depleted,additional amounts of the oil or small-molecule compounds can besupplied from the reservoir.

In an embodiment, the present invention provides a method for reducing,inhibiting, or modulating the behaviors of non-adherent pathogens inmedia surrounding a substrate comprising the steps of a) providing amultilayer film on the substrate, said multilayer film comprising:

i) one or more bilayers, wherein each bilayer comprises a first polymerlayer in contact with a second polymer layer, where said multilayerpolymer film has nanoscale or microscale porosity;

ii) an oil selected from the group consisting of a silicone oil, avegetable oil, a mineral oil, a thermotropic liquid crystal, andcombinations thereof, wherein said oil coats at least a portion of themultilayer polymer film and said oil at least partially fills the poresof at least a portion of said multilayer polymer film; and

-   -   iii) one or more small-molecule compounds able to reduce,        inhibit, or modulate the behaviors of said pathogens upon        contact with said pathogens, wherein the one or more        small-molecule compounds are located on the surface of said one        or more bilayers, within said oil, or both; and    -   b) controllably releasing an effective amount of the one or more        small-molecule compounds from said multilayer film into said        media, wherein the one or more small-molecule compounds contact        the pathogens thereby reducing the number of pathogens,        inhibiting the growth or colonization of the pathogens, or        modulating their virulence.

Preferably, the surface comprises one or more optionally functionalizedPVDMA/PEI bilayers and the oil is a silicone oil. Preferably, the one ormore small-molecule compounds comprise an antimicrobial agent, anantifungal agent, an antibacterial agent, an agent that modulatesbacterial or fungal quorum sensing, an agent that attenuates virulence,or a combination thereof, and the non-adherent pathogens comprisebacteria, fungi, or a combination thereof. Optionally, the one or moresmall-molecule compounds is a natural or synthetic antibiotic agent,natural or synthetic antifungal agent, a quorum sensing modulator or acombination thereof. In an embodiment, the one or more small-moleculecompounds is an AHL derivative or ABI derivative. In an embodiment, theone or more small-molecule compounds is selected from the groupconsisting of:

or combinations thereof.

Thus, the methods described herein can be used to fabricate physicallyand chemically durable SLIPS coatings on objects of arbitrary shape,size, and topology (e.g., on curved surfaces, insides of hollow tubes,etc.). Specifically these slippery surfaces could be used as antifoulingsurfaces, anti-bacterial/fungal surfaces where the liquid phases used toimpart anti-fouling properties can also be used as reservoirs for thecontrolled release of other active agents (e.g., antibiotics,antimicrobial agents, or anti-biofilm agents) that can reduce or inhibitnon-adherent pathogens in the surrounding media.

BRIEF DESCRIPTION OF THE DRAWINGS

FIG. 1. (A) Schematic illustration showing a side-view depiction of thebeading and sliding of an aqueous droplet on horizontally placed (top)and tilted (bottom) ‘slippery’ liquid-infused porous surfaces (SLIPS).(B-C) Digital pictures, acquired from a top down vantage point, of a 50μL droplet of C. albicans inoculum incubated on the surfaces of (B)SLIPS-coated glass substrates and (C) bare glass substrates for 3 hours;droplets are shown after staining with crystal violet and either beforeor after tilting from horizontal (at the angles indicated) and afterwashing with DI water; solid and dotted circles mark the originallocations of the droplets before sliding; arrows indicate direction ofsliding. (D-I) Bright-field (D,G) and fluorescence (E-F, H-I) microscopyimages of bare glass (D-F) and SLIPS-coated (G-I) glass substrates afterincubation with droplets of C. albicans inocula; droplets were incubatedfor 3 hours, stained with FUN-1 dye, and tilted to permit aqueous fluidsto slide away from the original location of the droplet prior toimaging; green fluorescence indicates cytoplasmic staining, red stainmarks intravacuolar structures in metabolically active (live) cells.Scale bars, including insets, are 100 μm.

FIG. 2: (A-L) Phase contrast and fluorescence microscopy images of bareglass (A-C), SLIPS-coated glass (D-F), oil-smeared glass (oil only;G-I), and porous polymer matrix (matrix only; J-L) surfaces immersed insuspensions of C. albicans for 24 hours; cells were stained with FUN-1fluorescent dye prior to imaging. (M) Plot showing the quantifiedmetabolic activity of C. albicans on the surfaces of bare glass,SLIPS-coated glass, oil-smeared glass (oil only), and porous polymermultilayers (matrix only) after immersion in suspensions of C. albicansfor 24 hours; metabolic activity was quantified using an XTT assay.Scale bars are 200 μm. Error bars represent standard deviation.

FIG. 3: (A-F) Bright-field microscopy images showing the surfaces ofbare glass and SLIPS-coated glass substrates after three consecutive24-hour challenges with C. albicans suspensions. (G-H) Fluorescencemicroscopy images of the same surfaces shown in (E-F) after stainingwith the FUN-1 fluorescent dye. (I) Plot showing the quantifiedmetabolic activity of C. albicans on the surfaces of bare glass andSLIPS-coated glass after the third consecutive challenge; metabolicactivity was quantified using an XTT assay. Scale bars are 100 μm. Errorbars represent standard deviation.

FIG. 4: (A-B) Lower- and higher-magnification SEM images showing thesurfaces of porous polymer multilayers fabricated on the inner surfacesof catheter tubes prior to the infusion of silicone oil; tube segmentswere sliced longitudinally prior to imaging (scale bar in A=100 μm;scale bar in B=2 μm). (C-E) Digital images showing the sliding ofaliquots of aqueous TMR (5 μL) inside SLIPS-coated catheter tubes tiltedat an angle of 10°. (F) Digital image showing an aliquot of aqueous TMRat the top end of a bare catheter tube. (G-H) Plots showing thequantified metabolic activity of C. albicans associated with thesurfaces (G) of bare and SLIPS-coated catheter tubes and in droplets ofyeast inocula (H) collected from bare and SLIPS-coated catheter tubesafter 4 hours of incubation. Error bars represent standard deviation.(I-N) Lower- and higher-magnification SEM images of bare (I-K) andSLIPS-coated (L-N) catheter tubes after inoculation with suspensions ofC. albicans for 4 hours; samples were prepared by conventionalcritical-point drying and tubes were sliced longitudinally prior toimaging. Scale bars in panels I,L; panels J,M and panels K,N are 200 μm,50 μm, and 10 μm, respectively.

FIG. 5: (A-F) Fluorescence microscopy images of the surfaces of bareglass and SLIPS-coated glass substrates after incubation in suspensionsof E. coli (A-B), P. aeruginosa (CD), and S. aureus (E-F) for 24 hours;samples were stained with a SYTO-9 green fluorescent nucleic acid stainprior to imaging. (G-H) Fluorescence microscopy images of bare glass andSLIPS-coated glass substrates after incubation with mammalian HeLa cellsfor 72 hours; samples were stained with calcein-AM prior to imaging.Scale bars are 100 μm.

FIG. 6: (A) Plot showing the advancing water contact angles, θadv, andcontact angle hystereses, θhys, of water droplets (8 μL) on SLIPS loadedwith triclosan (at levels ranging from 0 μg/cm² to 500 μg/cm²). (B)Digital pictures, acquired from a top-down perspective, showing thesliding of a droplet of aqueous TMR (15 μL) on a triclosan-loaded SLIP(loading=500 μg/cm²; tilt angle=10°). (C) Plot showing the release oftriclosan from triclosan-loaded SLIPS (closed circles; loading=500μg/cm²) upon incubation in PBS buffer at 37° C.; open circles correspondto the release profile of otherwise identical films not loaded withtriclosan; the inset of panel C shows the molecular structure oftriclosan. Error bars represent standard deviation.

FIG. 7: Plot showing the quantified metabolic activity of C. albicansassociated with the surfaces (left) of bare glass, SLIPS-coated glass,and triclosan-loaded SLIPS-coated glass and (right) the metabolicactivity of planktonic yeast growing in the surrounding medium;metabolic activities were quantified using an XTT assay; all data arenormalized with respect to the metabolic activities measured forexperiments using bare glass substrates. Substrates were incubated withC. albicans inoculum for 24 hours, removed from wells, and the surfacesand remaining cell suspensions were quantified separately. Error barsrepresent standard deviation; *indicates p<0.05 by a two-tailed T-test.

FIG. 8: (A) Plot showing the quantified metabolic activity of C.albicans associated with the surfaces of bare glass (black) andtriclosan-loaded SLIPS (grey) after each of five consecutive 24-hourchallenges in C. albicans inoculum, as determined using an XTT assay.(B) Plot showing the quantified metabolic activity of C. albicansassociated with the surfaces of bare glass and triclosan-loaded SLIPSafter continuous incubation in C. albicans inoculum for 7 days. Errorbars represent standard deviation.

FIG. 9: Plot showing the quantified metabolic activity of C. albicansassociated with the surfaces (left) of bare catheter tubes andtriclosan-loaded SLIPS-coated catheter tubes and (right) the metabolicactivity of planktonic cells growing in the surrounding intraluminalmedium; metabolic activities were quantified using an XTT assay; alldata are normalized with respect to the metabolic activities measuredfor experiments using bare catheter tubes. Substrates were incubatedwith C. albicans inoculum for 4 hours, and the surfaces and remainingcell suspensions were quantified separately. Error bars representstandard deviation.

FIG. 10: (A-D) Representative bright-field (A,C) and fluorescence (B,D)microscopy images showing the edges of SLIPS-coated glass substratesincubated in the presence of C. albicans for 24 hours. Samples werestained with FUN-1 fluorescent dye prior to imaging, and show small andisolated patches of biofilm (green) located along the edges and cornersof the coated substrates. The approximate locations of the edges of thesubstrates are indicated with black or white dotted lines. Scalebars=100 μm.

FIG. 11: (A-L) Representative fluorescence microscopy images ofsubstrates incubated in the presence of C. albicans for 24 hours.Samples were stained with FUN-1 fluorescent dye prior to imaging; greenindicates cytoplasmic staining and red indicates intravacuolarstructures in live cells. The images show results for both bare andSLIPS-coated glass substrates (A-D), plastic PET film substrates (E-H),and aluminum foil substrates (I-L). Scale bars are 100 μm. (M) Plotshowing the quantified metabolic activity of C. albicans on bare andSLIPS-coated glass, PET, and aluminum foil substrates after immersion insuspensions of C. albicans for 24 hours; metabolic activity wasquantified using an XTT assay. Error bars represent standard deviation.

FIG. 12: Representative bright-field (A,D) and fluorescence (B-C, E-F)microscopy images showing the inner surfaces of bare and SLIPS-coatedPTFE tubes incubated with C. albicans inocula for 4 hours. Samples werestained with FUN-1 fluorescent dye prior to imaging; green indicatescytoplasmic staining and red indicates intravacuolar structures in livecells. The textures observed in panel D arise from the features of thepolymer coating and, as indicated by the results in panels E-F, do notarise from the presence of biofilm. Scale bars are 100 μm.

FIG. 13: (A-P) Representative bright-field and fluorescence microscopyimages showing bare glass and SLIPS-coated glass substrates incubated inthe presence of E. coli, P. aeruginosa, S. aureus, or mammalian HeLacells. Scale bars are 100 μm. Films were incubated in the presence ofbacteria (A-L) for 24 hours and then treated with SYTO-9 greenfluorescent nucleic acid stain prior to imaging. Films were incubated inthe presence of mammalian cells (M-P) for 72 hours and stained withcalcein-AM prior to imaging.

FIG. 14: Digital photographs providing a visual indication of relativelevels of metabolic activity exhibited by C. albicans biofilms formed inthe presence of bare glass (top) or triclosan-loaded, SLIPS-coated glasssubstrates (as determined by an XTT assay; three replicates are shown).The intensity of the orange color (shown here in grey scale) indicatesrelative levels of metabolic activity.

FIG. 15: Schematic illustration showing the fabrication of thecontrolled release SLIPS used in one embodiment of the invention. (A)Reactive and nanoporous polymer multilayers (grey) are functionalizedwith n-decylamine to render them hydrophobic. (B) Small molecules(diamonds) are loaded into porous multilayers by adding an acetonesolution of the agents to dried films and allowing the solvent toevaporate. (C) Silicone oil is infused into the multilayers. (D) Thesecontrolled release SLIPS gradually release loaded compounds into aqueoussolution when hosted in aqueous environments; these dual-action SLIPSprevent the colonization of bacteria directly on the coated surface andcan also modulate the behaviors of planktonic bacteria.

FIG. 16: Structures of the small molecule anti-virulence agents used inone embodiment of the invention.

FIG. 17: Plot showing percent release versus time for the release ofcompounds E22 (triangles), C14 (diamonds), and DMABI (squares) fromSLIPS-coated surfaces incubated in PBS buffer at 37° C. The inset showsthe same results focusing specifically on the first 50 hours of release.Data points represent the mean of four replicates. The percentage ofeach compound released was calculated based on the total amount ofcompound initially loaded.

FIG. 18: Inhibition of pyocyanin production by QSIs released fromSLIPS-coated surfaces. P. aeruginosa cultures were grown in the presenceof SLIPS loaded with the indicated compounds and the final amount ofpyocyanin in the culture was quantified after 17 hours of incubation.Compounds were loaded at levels estimated to give a final releasedconcentrations of 100 μM (for experiments involving a single loadedcompound) or 50 μM (for experiments involving two loaded compounds).Error bars represent the standard error of three independent replicates(n=3). ***=p<0.0005.

FIG. 19: SLIPS loaded with DMABI resist fouling on the substrate surfaceand inhibit biofilm formation on surrounding uncoated surfaces. Glass orSLIPS-coated substrates were submerged in P. aeruginosa cultures at thebottoms of the wells of 12-well microtiter plates and incubated for 24h. Substrates were then removed and the attached biofilms were stainedwith either SYTO 9 or CV. (A) Representative fluorescence microscopyimages of P. aeruginosa biofilm near to the center of glass (left) orSLIPS-coated (right) substrates. (B) Representative image of CV-stainedbiofilms attached to glass substrates, SLIPS, or SLIPS loaded withDMABI. (C) Representative image of CV staining of the bottoms of thewells of the 12-well microtiter plate (after the removal of thesubstrates; shown in panel A). (D) Quantification of biofilm formed onthe glass and SLIPS substrates shown in panel B. (E) Quantification ofbiofilm formed on the surrounding well bottoms shown in panel C, showinga reduction of biofilm in wells that contained DMABI-loaded SLIPS. (F)Representative fluorescence microscopy images of biofilm formed on theexposed glass edges of SLIPS substrates without DMABI (left) and withloaded DMABI (right). Scale bars in A and F represent 400 μm. Error barsin D and E represent the standard error of three independent experiments(n=3). *=p<0.05. NS=no statistical difference.

FIG. 20A and FIG. 20B: Plot showing a release curve (determined usingfluorescence intensity) of a fluorophore (tetramethyl rhodamine; TMR)which was loaded into SLIPS by loading TMR into silicone oil and theninfusing the TMR/oil solution into the polymer matrix (FIG. 20A). Plotshowing a release curve of TMR which was loaded into SLIPS by firstloading TMR into the matrix followed by infusing the matrix with thesilicone oil (FIG. 20B).

DETAILED DESCRIPTION OF THE INVENTION Definitions

As used herein, the term “slippery” refers to surfaces that allow liquiddroplets to slide off the surface with sliding angles of 10° or less,preferably 5° or less, 2.5° or less, or 2° or less.

As used herein, the term “controllably released” refers to a molecule,drug and/or compound which is initially contained within the porousmatrix and/or oil of a slippery liquid-infused porous surface (SLIPS)and is progressively released into the surrounding media over aconsistent period of time. In some embodiments, the time require torelease at least 50% of the molecule, drug and/or compound into thesurrounding media is 6 hours or more, preferably 24 hours or more, 4days or more, preferably 10 days or more, 20 days or more, 30 days ormore, 60 days or more, 100 days or more, 120 days or more, or 180 daysor more.

As used herein, “functionalized polymer” refers to a polymer in which atleast a portion of the individual monomer units are substituted with aspecific functional group. For the functionalized polymers of thepresent invention, at least 1% or more, at least 2% or more, at least 5%or more, at least 10% or more, at least 15% or more, at least 20% ormore, at least 30% or more, at least 50% or more, at least 75% or more,or at least 90% or more of the portion of the monomer units issubstituted with a specific functional group.

An “amine reactive group” or “hydroxyl reactive group” can be anyfunctional group able to react with an amine group or hydroxyl group,respectively.

As used herein, the term “anti-fouling” refers to a material's abilityto resist adhesion by an undesirable material, such as oils, organiccompounds, and organisms. In particular, it is desirable to prevent orreduce the adhesion of hydrophobic compounds and organisms to a materialwhich is submerged or in contact with water.

The term “nanoscale” refers to a length less than 1,000 nm, preferablyless than 100 nm, and the term “microscale” refers to a length less than1,000 μm, preferably less than 100 μm.

The term “alkyl” refers to a monoradical of a branched or unbranched(straight-chain or linear) saturated hydrocarbon and to cycloalkylgroups having one or more rings. Alkyl groups as used herein includethose having from 1 to 20 carbon atoms, preferably having from 1 to 12carbon atoms. Alkyl groups include small alkyl groups having 1 to 3carbon atoms. Alkyl groups include medium length alkyl groups havingfrom 4-10 carbon atoms. Alkyl groups include long alkyl groups havingmore than 10 carbon atoms, particularly those having 10-20 carbon atoms.Cycoalkyl groups include those having one or more rings. Cyclic alkylgroups include those having a 3-, 4-, 5-, 6-, 7-, 8-, 9-, 10-, 11- or12-member carbon ring and particularly those having a 3-, 4-, 5-, 6-, or7-member ring. The carbon rings in cyclic alkyl groups can also carryalkyl groups. Cyclic alkyl groups can include bicyclic and tricyclicalkyl groups. Alkyl groups are optionally substituted. Substituted alkylgroups include among others those which are substituted with arylgroups, which in turn can be optionally substituted. Specific alkylgroups include methyl, ethyl, n-propyl, iso-propyl, cyclopropyl,n-butyl, s-butyl, t-butyl, cyclobutyl, n-pentyl, branched-pentyl,cyclopentyl, n-hexyl, branched hexyl, and cyclohexyl groups, all ofwhich are optionally substituted. Substituted alkyl groups include fullyhalogenated or semihalogenated alkyl groups, such as alkyl groups havingone or more hydrogens replaced with one or more fluorine atoms, chlorineatoms, bromine atoms and/or iodine atoms. Substituted alkyl groupsinclude fully fluorinated or semifluorinated alkyl groups, such as alkylgroups having one or more hydrogens replaced with one or more fluorineatoms. An alkoxy group is an alkyl group linked to oxygen and can berepresented by the formula R—O. Examples of alkoxy groups include, butare not limited to, methoxy, ethoxy, propoxy, butoxy and heptoxy. Alkoxygroups include substituted alkoxy groups wherein the alky portion of thegroups is substituted as provided herein in connection with thedescription of alkyl groups.

The term “alkenyl” refers to a monoradical of a branched or unbranchedunsaturated hydrocarbon group having one or more double bonds and tocycloalkenyl groups having one or more rings wherein at least one ringcontains a double bond. Alkenyl groups include those having 1, 2 or moredouble bonds and those in which two or more of the double bonds areconjugated double bonds. Alkenyl groups include those having from 2 to20 carbon atoms, preferably having from 2 to 12 carbon atoms. Alkenylgroups include small alkenyl groups having 2 to 3 carbon atoms. Alkenylgroups include medium length alkenyl groups having from 4-10 carbonatoms. Alkenyl groups include long alkenyl groups having more than 10carbon atoms, particularly those having 10-20 carbon atoms. Cycloalkenylgroups include those having one or more rings. Cyclic alkenyl groupsinclude those in which a double bond is in the ring or in an alkenylgroup attached to a ring. Cyclic alkenyl groups include those having a3-, 4-, 5-, 6-, 7-, 8-, 9-, 10-, 11- or 12-member carbon ring andparticularly those having a 3-, 4-, 5-, 6- or 7-member ring. The carbonrings in cyclic alkenyl groups can also carry alkyl groups. Cyclicalkenyl groups can include bicyclic and tricyclic alkyl groups. Alkenylgroups are optionally substituted. Substituted alkenyl groups includeamong others those which are substituted with alkyl or aryl groups,which groups in turn can be optionally substituted. Specific alkenylgroups include ethenyl, prop-1-enyl, prop-2-enyl, cycloprop-1-enyl,but-1-enyl, but-2-enyl, cyclobut-1-enyl, cyclobut-2-enyl, pent-1-enyl,pent-2-enyl, branched pentenyl, cyclopent-1-enyl, hex-1-enyl, branchedhexenyl, cyclohexenyl, all of which are optionally substituted.Substituted alkenyl groups include fully halogenated or semihalogenatedalkenyl groups, such as alkenyl groups having one or more hydrogensreplaced with one or more fluorine atoms, chlorine atoms, bromine atomsand/or iodine atoms. Substituted alkenyl groups include fullyfluorinated or semifluorinated alkenyl groups, such as alkenyl groupshaving one or more hydrogens replaced with one or more fluorine atoms.

The term “aryl” refers to a chemical group having one or more 5-, 6- or7-member aromatic or heterocyclic aromatic rings. An aromatichydrocarbon is a hydrocarbon with a conjugated cyclic molecularstructure. Aryl groups include those having from 4 to 30 carbon atoms,preferably having from 6 to 18 carbon atoms. Aryl groups can contain asingle ring (e.g., phenyl), one or more rings (e.g., biphenyl) ormultiple condensed (fused) rings, wherein at least one ring is aromatic(e.g., naphthyl, dihydrophenanthrenyl, fluorenyl, or anthryl).Heterocyclic aromatic rings can include one or more N, O, or S atoms inthe ring. Heterocyclic aromatic rings can include those with one, two orthree N, those with one or two O, and those with one or two S, orcombinations of one or two or three N, O or S. Aryl groups areoptionally substituted. Substituted aryl groups include among othersthose which are substituted with alkyl or alkenyl groups, which groupsin turn can be optionally substituted. Specific aryl groups includephenyl groups, biphenyl groups, pyridinyl groups, and naphthyl groups,all of which are optionally substituted. Substituted aryl groups includefully halogenated or semihalogenated aryl groups, such as aryl groupshaving one or more hydrogens replaced with one or more fluorine atoms,chlorine atoms, bromine atoms and/or iodine atoms. Substituted arylgroups include fully fluorinated or semifluorinated aryl groups, such asaryl groups having one or more hydrogens replaced with one or morefluorine atoms. Aryl groups include, but are not limited to, aromaticgroup-containing or heterocylic aromatic group-containing groupscorresponding to any one of the following benzene, naphthalene,naphthoquinone, diphenylmethane, fluorene, fluoranthene, anthracene,anthraquinone, phenanthrene, tetracene, naphthacenedione, pyridine,quinoline, isoquinoline, indoles, isoindole, pyrrole, imidazole,oxazole, thiazole, pyrazole, pyrazine, pyrimidine, purine,benzimidazole, furans, benzofuran, dibenzofuran, carbazole, acridine,acridone, phenanthridine, thiophene, benzothiophene, dibenzothiophene,xanthene, xanthone, flavone, coumarin, azulene or anthracycline. As usedherein, a group corresponding to the groups listed above expresslyincludes an aromatic or heterocyclic aromatic radical, includingmonovalent, divalent and polyvalent radicals, of the aromatic andheterocyclic aromatic groups listed above provided in a covalentlybonded configuration in the compounds of the present invention. Arylgroups optionally have one or more aromatic rings or heterocyclicaromatic rings having one or more electron donating groups, electronwithdrawing groups and/or targeting ligands provided as substituents.

Arylalkyl groups are alkyl groups substituted with one or more arylgroups wherein the alkyl groups optionally carry additional substituentsand the aryl groups are optionally substituted. Specific alkylarylgroups are phenyl-substituted alkyl groups, e.g., phenylmethyl groups.Alkylaryl groups are alternatively described as aryl groups substitutedwith one or more alkyl groups wherein the alkyl groups optionally carryadditional substituents and the aryl groups are optionally substituted.Specific alkylaryl groups are alkyl-substituted phenyl groups such asmethylphenyl. Substituted arylalkyl groups include fully halogenated orsemihalogenated arylalkyl groups, such as arylalkyl groups having one ormore alkyl and/or aryl having one or more hydrogens replaced with one ormore fluorine atoms, chlorine atoms, bromine atoms and/or iodine atoms.

Optional substitution of any alkyl, alkenyl and aryl groups includessubstitution with one or more of the following substituents: halogens,—CN, —COOR, —OR, —COR, —OCOOR, —CON(R)₂, —OCON(R)₂, —N(R)₂, —NO₂, —SR,—SO₂R, —SO₂N(R)₂ or —SOR groups. Optional substitution of alkyl groupsincludes substitution with one or more alkenyl groups, aryl groups orboth, wherein the alkenyl groups or aryl groups are optionallysubstituted. Optional substitution of alkenyl groups includessubstitution with one or more alkyl groups, aryl groups, or both,wherein the alkyl groups or aryl groups are optionally substituted.Optional substitution of aryl groups includes substitution of the arylring with one or more alkyl groups, alkenyl groups, or both, wherein thealkyl groups or alkenyl groups are optionally substituted.

Optional substituents for alkyl and alkenyl groups include among others:

-   -   —COOR where R is a hydrogen or an alkyl group or an aryl group        and more specifically where R is methyl, ethyl, propyl, butyl,        or phenyl groups all of which are optionally substituted;    -   —COR where R is a hydrogen, or an alkyl group or an aryl groups        and more specifically where R is methyl, ethyl, propyl, butyl,        or phenyl groups all of which groups are optionally substituted;    -   —CON(R)₂ where each R, independently of each other R, is a        hydrogen or an alkyl group or an aryl group and more        specifically where R is methyl, ethyl, propyl, butyl, or phenyl        groups all of which groups are optionally substituted; R and R        can form a ring which may contain one or more double bonds;    -   —OCON(R)₂ where each R, independently of each other R, is a        hydrogen or an alkyl group or an aryl group and more        specifically where R is methyl, ethyl, propyl, butyl, or phenyl        groups all of which groups are optionally substituted; R and R        can form a ring which may contain one or more double bonds;    -   —N(R)₂ where each R, independently of each other R, is an alkyl        group, acyl group or an aryl group and more specifically where R        is methyl, ethyl, propyl, butyl, or phenyl or acetyl groups all        of which are optionally substituted; or R and R can form a ring        which may contain one or more double bonds.    -   —SR, —SO₂R, or —SOR where R is an alkyl group or an aryl groups        and more specifically where R is methyl, ethyl, propyl, butyl,        phenyl groups all of which are optionally substituted; for —SR,        R can be hydrogen;    -   —OCOOR where R is an alkyl group or an aryl groups;    -   —SO₂N(R)₂ where R is a hydrogen, an alkyl group, or an aryl        group and R and R can form a ring;    -   —OR where R is H, alkyl, aryl, or acyl; for example, R can be an        acyl yielding —OCOR* where R* is a hydrogen or an alkyl group or        an aryl group and more specifically where R* is methyl, ethyl,        propyl, butyl, or phenyl groups all of which groups are        optionally substituted.

As used herein, the term “alkylene” refers to a divalent radical derivedfrom an alkyl group or as defined herein. Alkylene groups in someembodiments function as attaching and/or spacer groups in the presentcompositions. Compounds of the present invention include substituted andunsubstituted C₁-C₃₀ alkylene, C₁-C₁₂ alkylene and C₁-C₅ alkylenegroups. The term “alkylene” includes cycloalkylene and non-cyclicalkylene groups.

As used herein, the term “cycloalkylene” refers to a divalent radicalderived from a cycloalkyl group as defined herein. Cycloalkylene groupsin some embodiments function as attaching and/or spacer groups in thepresent compositions. Compounds of the present invention includesubstituted and unsubstituted C₁-C₃₀ cycloalkenylene, C₁-C₁₂cycloalkenylene and C₁-C₅ cycloalkenylene groups.

As used herein, the term “alkenylene” refers to a divalent radicalderived from an alkenyl group as defined herein. Alkenylene groups insome embodiments function as attaching and/or spacer groups in thepresent compositions. Compounds of the present invention includesubstituted and unsubstituted C₁-C₂₀ alkenylene, C₁-C₁₂ alkenylene andC₁-C₅ alkenylene groups. The term “alkenylene” includes cycloalkenyleneand non-cyclic alkenylene groups.

As used herein, the term “cycloalkenylene” refers to a divalent radicalderived from a cylcoalkenyl group as defined herein. Cycloalkenylenegroups in some embodiments function as attaching and/or spacer groups inthe present compositions.

Specific substituted alkyl groups include haloalkyl groups, particularlytrihalomethyl groups and specifically trifluoromethyl groups. Specificsubstituted aryl groups include mono-, di-, tri, tetra- andpentahalo-substituted phenyl groups; mono-, di-, tri-, tetra-, penta-,hexa-, and hepta-halo-substituted naphthalene groups; 3- or4-halo-substituted phenyl groups, 3- or 4-alkyl-substituted phenylgroups, 3- or 4-alkoxy-substituted phenyl groups, 3- or4-RCO-substituted phenyl, 5- or 6-halo-substituted naphthalene groups.More specifically, substituted aryl groups include acetylphenyl groups,particularly 4-acetylphenyl groups; fluorophenyl groups, particularly3-fluorophenyl and 4-fluorophenyl groups; chlorophenyl groups,particularly 3-chlorophenyl and 4-chlorophenyl groups; methylphenylgroups, particularly 4-methylphenyl groups, and methoxyphenyl groups,particularly 4-methoxyphenyl groups.

As used herein, the term “halo” refers to a halogen group such as afluoro (—F), chloro (—Cl), bromo (—Br) or iodo (—I).

As to any of the above groups which contain one or more substituents, itis understood, that such groups do not contain any substitution orsubstitution patterns which are sterically impractical and/orsynthetically non-feasible. In addition, the compounds of this inventioninclude all stereochemical isomers arising from the substitution ofthese compounds.

Overview

A Controlled Release Approach for Utilizing SLIPS to Prevent MicrobialSurface Fouling and Kill Non-Adherent Pathogens in Surrounding Media.Surface-associated fouling by bacteria is a common and persistentchallenge facing the use of biomedical devices, industrial equipment,and many consumer products. The development of strategies that can slowor prevent microbial attachment and attenuate other bacterial behaviorson surfaces is thus an important element in the design of materials andcoatings intended for use in wet environments. Embodiments of thepresent invention are motivated broadly by the recent development ofslippery liquid-infused porous surfaces (or ‘SLIPS’), which havephysical properties and behaviors that render them well suited for thedesign of anti-biofouling surfaces.

SLIPS comprise a porous or textured surface infused with a viscousliquid (e.g., perfluorinated lubricants, silicone oil, etc.). Thisgeneral design maintains the infused liquid as a thin, dynamic film atthe surface, creating a hydrophobic or omniphobic interface that allowsother fluids and substances to easily slide or ‘slip’ off the surfacewith sliding angles as low as 2°. Several recent reports reveal SLIPS tobe a promising platform for the development of new anti-biofoulinginterfaces for biological and environmental applications. Indeed, SLIPShave been reported to resist fouling by a broad range of organisms,including clinically important bacterial and fungal pathogens, marinebarnacle cyprids, and mammalian cells.

Slippery character is the sine qua non of a SLIPS-coated surface, butthis essential quality only allows SLIPS to prevent fouling by organismson the surfaces to which these coatings are physically applied.Conventional SLIPS-coated surfaces, for example, cannot prevent bacteriafrom colonizing other nearby (non-SLIPS-coated) surfaces. ConventionalSLIPS also do not kill bacteria—organisms that are prevented fromadhering to SLIPS-coated surfaces remain alive in the surroundingmedium, and SLIPS do not currently have inherent mechanisms throughwhich they can prevent these non-adherent (or ‘planktonic’) bacteriafrom producing toxins or engaging in other virulent behaviors, includingforming biofilms on nearby unprotected surfaces.

To address these issues and develop new slippery anti-fouling surfacesthat can also exert control over the behaviors of microorganisms insurrounding media, the present invention provides a new controlledrelease-based approach to the design of multifunctional SLIPS thatprevent biofouling by pathogenic fungal and bacterial cells and killplanktonic microorganisms in surrounding media. In this approach, theproperties of a porous polymer matrix and an infused silicone oil phaseare leveraged to sustain the long-term release of small-moleculecompounds, particularly agents directed toward microorganisms (such asbacteria and fungi). Experimental studies demonstrated that suchsmall-molecule anti-microbial agents can be readily incorporated intoSLIPS without impacting the anti-fouling properties of the SLIPSsurfaces, and that the slow release of such anti-microbial agents cankill planktonic fungal cells effectively and improve the overallantifouling and antifungal properties.

Such anti-microbial agents include, but are not limited to triclosan andother broad-spectrum antibiotics. It should be noted, however, that theuse of triclosan and other cytotoxic drugs (e.g., antibiotics) haveseveral disadvantages in applied contexts, including the fact that thewidespread use of these agents has led to evolved resistance in manyclinically relevant pathogens.

EXAMPLES Example 1—Fabrication of SLIPS Material with an Antimicrobial(Triclosan)

Many types of slippery liquid-infused porous surfaces (or ‘SLIPS’) canresist adhesion and colonization by microorganisms. These ‘slippery’materials thus offer new approaches to prevent fouling on a range ofcommercial and industrial surfaces, including biomedical devices.However, while SLIPS can prevent fouling on surfaces to which they areapplied, they can currently do little to prevent the proliferation ofnon-adherent (planktonic) organisms, stop them from colonizing othersurfaces, or prevent them from engaging in other behaviors that couldlead to infection and associated burdens. The present examples providean approach to the design of multi-functional SLIPS that addresses theseissues and expands the potential utility of slippery surfaces inantimicrobial contexts.

This approach is based on the incorporation and controlled release ofsmall-molecule antimicrobial agents from the porous matrices used tohost infused slippery oil phases. The below examples demonstrate thatSLIPS fabricated using nanoporous polymer multilayers can prevent short-and longer-term colonization and biofilm formation by four common fungaland bacterial pathogens (Candida albicans, Pseudomonas aeruginosa,Escherichia coli, and Staphylococcus aureus), and that the polymer andoil phases comprising these materials can be exploited to load andsustain the release of triclosan, a model hydrophobic and broad-spectrumantimicrobial agent, into surrounding media. This approach both improvesthe inherent anti-fouling properties of these materials and endows themwith the ability to efficiently kill planktonic pathogens.

Finally, this approach can be used to fabricate dual-action SLIPS oncomplex surfaces, including the luminal surfaces of flexible cathetertubes. This strategy has the potential to be generally applicable and itis anticipated that the materials, strategies, and concepts reportedhere will enable new approaches to the design of slippery surfaces withimproved anti-fouling properties and open the door to new applicationsof slippery liquid-infused materials that host or promote the release ofa variety of other active agents.

Materials and Methods. In an embodiment,poly(2-vinyl-4,4-dimethylazlactone) (PVDMA) was synthesized byfree-radical polymerization of VDMA as described previously (Buck etal., Chem. Mater. 2010, 22, 6319). Branched poly(ethyleneimine) (BPEI;MW ˜25,000), propylamine, n-decylamine (95%), acetone (HPLC grade),tetrahydrofuran (THF, HPLC grade), and silicone oil for melting pointand boiling point apparatuses (viscosity=45-55 cSt at 25° C.) werepurchased from Sigma Aldrich (Milwaukee, Wis.). D-Glucamine (>95%) waspurchased from TCI America (Portland, Oreg.). Glass microscope slideswere purchased from Fischer Scientific (Pittsburgh, Pa.). Triclosan(5-chloro-2-(2,4-dichlorophenoxy)phenol) was purchased from Alfa Aesar(Ward Hill, Mass.).

Thin sheets of poly(ethylene terephthalate) film (PET; 0.004 in. thick)were purchased from McMaster Carr (Elmhurst, Ill.). Aluminum foil wasobtained from Reynolds Consumer Products, LLC (Richmond, Va.).Polyethylene tubing (PE-100, inner diameter=0.034 in.) was purchasedfrom Intramedic (Franklin Lakes, N.J.). Roswell Park Memorial Institute(RPMI) 1640 powder (with L-glutamine and phenol red, without HEPES andsodium bicarbonate), FUN-1 cell stain, fetal bovine serum (FBS) and2,3-bis-(2-methoxy-4-nitro-5-sulfophenyl)-2H-tetrazolium-5-carboxanilide(XTT) were purchased from Invitrogen (Grand Island, N.Y.).3-(N-Morpholino)propanesulfonic acid (MOPS), Tris-base,phosphate-buffered saline (PBS) liquid concentrate (10×), and NaCl werepurchased from Fisher Scientific (Pittsburgh, Pa.). Menadione,glutaraldehyde, and formaldehyde were purchased from Sigma (St. Louis,Mo.). Tween-20 was purchased from Acros (Grand Island, N.Y.). Osmiumtetroxide (4%) was purchased from Electron Microscopy Sciences(Hatfield, Pa.).

Escherichia coli strain ATCC 8739, Staphylococcus aureus strainATCC10390, and HeLa cells were purchased from American Type CultureCollection (ATTC; Manassas, Va.). Pseudomonas aeruginosa strain (PAO1)was obtained from the University of Rochester. SYTO-9 staining kits werepurchased from ThermoFisher (Waltham, Mass.). All chemicals, reagents,and solvents were used as received without further purification unlessotherwise noted.

General Considerations. Compressed air used to dry samples was filteredthrough a 0.2 μm membrane syringe filter. Scanning electron micrographswere acquired using a LEO 1530 scanning electron microscope (SEM) at anaccelerating voltage of 5 kV. Samples were coated with a thin layer ofgold using a gold sputterer operating at 45 mA under a vacuum pressureof 50 mTorr for 40 seconds prior to imaging. Critical point drying wasused to prepare fungal biofilm samples for SEM imaging using a CriticalPoint Dryer (Tousimis Samdri-815). Digital pictures were acquired usinga Canon PowerShot SX130 IS digital camera.

Water contact angles were measured using a Dataphysics OCA 15 Pluscontact angle goniometer at ambient temperature with 10 μL of deionizedwater. Fluorescence microscopy was performed using an Olympus IX71microscope and images were obtained using the MetaMorph Advanced version7.8.1.0 software package. Images were processed using NIH Image Jsoftware and Microsoft Powerpoint 2010. In all tube-based experimentsdescribed below in this example, the ends of the tubes were left open.In droplet-based and tube-based experiments, substrates were placed in ahumidified microenvironment before incubating at 37° C.

Absorbance measurements used in XTT assays were acquired at 490 nm usinga plate reader (EL800 Universal Microplate Reader, Bio-Tek Instruments,Inc.). The M9 buffer used in triclosan release experiments was preparedwith the following previously reported composition: 8.6 mM NaCl, 47.7 mMNa2HPO4, 21.7 mM KH2PO4, 18.7 mM NH₄Cl, pH 7.35 (see De Kievit et al.,Appl. Environ. Microbiol., 2001, 67, 1865; and Frei et al., Angew. Chem.Int. Ed. 2012, 51, 5226).

Covalent Layer-by-Layer Assembly of Porous Polymer Multilayers. PorousPEI/PVDMA multilayers were deposited on glass substrates using apreviously reported procedure (see Manna et al., Adv. Mater. 2015, 27,3007; and Manna et al., Adv. Funct. Mater. 2015, 25, 1672). Briefly: (i)substrates were submerged in a solution of PEI (20 mM in acetone withrespect to the polymer repeat unit) for 20 seconds; (ii) substrates wereremoved and immersed in an initial acetone bath for 20 seconds followedby a second acetone bath for 20 seconds; (iii) substrates were submergedin a solution of PVDMA (20 mM in acetone with respect to the polymerrepeat unit) for 20 seconds; and (iv) substrates were removed and rinsedagain using the procedure outlined under step (ii). This cycle wasrepeated 35 times to fabricate porous polymer multilayers consisting of35 PEI/PVDMA layer pairs (or ‘bilayers’).

The concentrations of the polymer solutions were maintained duringassembly by the addition of acetone as needed to compensate for solventevaporation after every dipping cycle. All other substrates used in thisstudy (e.g., catheter tubes, aluminum foil, PET film, and PTFE tubes)were also coated using this general protocol. Multilayers werecharacterized and used in subsequent experiments immediately or driedunder a stream of filtered, compressed air and stored in a vacuumdesiccator until use. All films were fabricated at ambient roomtemperature.

Chemical Functionalization of Multilayers and Infusion of Oils. Porouspolymer multilayers containing unreacted azlactone groups, prepared asdescribed above, were functionalized by treatment with solutions ofdecylamine (20 mM in THF) (see Manna et al., Adv. Mater. 2015, 27, 3007;and Manna et al., Adv. Mater. 2012, 24, 4291).

Functionalized films were then rinsed with THF and acetone and driedwith filtered air. The resulting hydrophobic multilayers were infusedwith lubricating liquids (e.g., silicone oil) using the followinggeneral protocols. For planar substrates, a droplet of 3 μL of oil wasplaced onto a film and spread over the surface using weighing paper. Forexperiments involving coated tubes, film-coated tubes were infused withoil by placing a 4 μL silicone oil droplet at the top end of a verticaltube and allowing gravity-driven spreading of the oil from the top tothe bottom of the tube. Excess oil was removed by tapping the bottom endof the tube on a disposable wipe.

Loading and Release of Triclosan. The small-molecule antimicrobial agenttriclosan was loaded into porous multilayers prior to the infusion ofsilicone oil by treating film-coated substrates with a desired number of10 μL droplets of a solution of triclosan (50 mg/mL) in acetone andallowing the acetone to evaporate under ambient conditions for 5minutes. All films were then dried under vacuum and infused withsilicone oil as described above. Characterization of the release oftriclosan from these loaded SLIPS was performed by incubating loadedSLIPS-coated substrates in 750 μL of PBS buffer at 37° C. The release oftriclosan as a function of time was monitored by UV/visspectrophotometry at a wavelength of 222 nm. At desired time points,buffer was removed for analysis using a pipette and replaced by a fresh750 μL aliquot.

Characterization of Fungal Cell Adhesion on SLIPS-Coated Substrates. C.albicans SC5314 cells were grown overnight at 30° C. in liquid yeastextract-peptone-dextrose (YPD) medium. Cells were washed with PBS andre-suspended in RPMI 1640 medium buffered with MOPS (pH 7.0). The celldensity was adjusted to 10⁷ cfu/mL with RPMI 1640 for experiments withSLIPS that did not contain triclosan, and to 10⁶ cfu/mL for experimentsusing SLIPS loaded with triclosan. For initial screens, 50 μL dropletsof cell suspension were placed on both planar SLIPS-coated and planarbare glass surfaces and incubated at 37° C.

Substrates were removed from the incubator after 3 hours andcharacterized using either a macroscopic crystal violet stain or amicroscopic yeast cell viability dye. The macroscopic cell stain assaywas performed by adding 20 μL of a crystal violet solution [CV; 1% (w/w)in deionized water] to droplets of yeast sitting on the surfaces ofSLIPS-coated glass or bare glass substrates and incubating at 37° C. for30 minutes. Substrates were then removed from the incubator and placedin an inclined position at a desired angle to characterize and comparethe anti-fouling properties of the substrates. Microscopic cellviability assays were performed by adding 25 μL of a 10 μM solution ofthe fungal cell viability probe FUN-1 to the yeast droplets andincubating at 37° C. for at least 30 minutes before imaging using afluorescence microscope.

Estimation of Biofilm Adhesion on SLIPS-Coated Substrates. C. albicansSC5314 cells were grown overnight at 30° C. in liquid YPD medium. Cellswere washed with PBS and resuspended in RPMI 1640 medium buffered withMOPS (pH 7.0) and supplemented with 5% fetal bovine serum to stimulatebiofilm formation. The cell density was manually adjusted to 10⁶ cfu/mL.SLIPS-coated substrates and other control substrates (bare glass,silicone oil-wetted glass, and multilayer-coated glass without siliconeoil) were individually submerged in 1 mL of cell suspension (10⁶ cfu/mL)in each well of a 24-well plate. The plate was incubated at 37° C. for24 hours, after which time either (i) the biofilms formed on thesurfaces of the substrates were visualized using the cell stain FUN-1(200 μL of 10 μM FUN-1, incubated for 30 minutes at 37° C.) or (ii) themetabolic activity of the cells was evaluated. For evaluation ofmetabolic activity, substrates were carefully removed from the wells,excess liquid was removed and substrates were placed in new wells; 200μL of XTT solution (0.5 g/L in PBS, pH 7.4, containing 3 μM menadione inacetone) was added to every well containing a substrate, includingnegative control wells that did not have any yeast or substrate in them.

After 2 hours of incubation at 37° C., 75 μL of the supernatant wastransferred to a 96-well plate and the absorbance of the solution at 490nm was measured. Background absorbance from wells containing medium andXTT alone was subtracted from all readings and results were plottedrelative to the absorbance values of wells containing samples ofsolution from control untreated surfaces. Similar sets of experimentswere carried out for SLIPS-coated PET film and SLIPS-coated aluminumfoil. For studies involving the use of triclosan-loaded SLIPS, themetabolic activity of cells in solution was also quantified (in bothtriclosan-loaded and non-triclosan-loaded control coatings) in additionto characterization of the metabolic activity of surface-associatedcells.

For multiple challenge experiments, substrates were incubated withyeast, as described above. At the end of each 24-hour period, substrateswere removed from their wells and imaged to characterize the extent towhich yeast was adhered on the surface. The SLIPS-coated substrates werethen incubated in fresh C. albicans suspensions to perform the nextchallenge (new bare glass substrates were used in control experiments).Three such 24-hour challenges were performed, and at the end of thethird challenge, XTT was used to quantify the reduction in metabolicactivity of cells on the surface of the SLIPS substrates compared toglass controls. Similar multiple challenge experiments, consisting offive consecutive challenges, were performed using triclosan-loadedSLIPS. In those experiments, the metabolic activities of bothsubstrate-associated and planktonic cells were quantified at the end ofevery challenge.

Characterization of Fungal Cell Adhesion in SLIPS-Coated Catheter Tubes.A 15 μL aliquot of a C. albicans suspension (10⁷ cfu/mL) was incubatedin 3 cm segments of SLIPS-coated PE tubes at 37° C. After 4 hours, theamount of attached cells in the tubes was both qualitatively andquantitatively estimated using three different methods: (i) a metabolicXTT assay, (ii) by visualizing the biofilm formed using SEM, and (iii)using the FUN-1 cell stain assay as described above. For the metabolicassay, yeast solution was removed from the tubes and placed in a wellcontaining 50 μL of XTT (0.5 g/L in PBS, pH 7.4, containing 3 μMmenadione in acetone).

A 15 μL aliquot of XTT was also incubated inside the emptied tubes. Bothpreparations were incubated at 37° C. for 2 hours, after which theabsorbance of the solutions was measured at 490 nm to quantify therelative metabolic activity of yeast on the surface of the tubes andinside the tubes. For analysis by SEM, catheter tube segments wereplaced in a fixative solution [1% (v/v) glutaraldehyde and 4% (v/v)formaldehyde] overnight at 4° C. The samples were rinsed in PBS (0.1 M)for 10 minutes and then placed in osmium tetroxide (1%) for 30 minutes,followed by 10 minutes in PBS (0.1 M). The samples were subsequentlydehydrated in a series of ethanol washes (30%, 50%, 70%, 85%, 95%, and100% for 20 minutes each). Final desiccation was accomplished bycritical point drying.

Specimens were mounted on aluminum stubs and sliced open to reveal theinside of the catheter, then sputter coated with gold-palladium. Sampleswere then imaged in high-vacuum mode at 5 kV. The antifungal andanti-biofilm activities of triclosan-loaded, SLIPS-coated PTFE tubeswere characterized by incubating 40 μL of a 10⁶ cfu/mL C. albicans cellsuspension in a 2 cm tube segment for 4 hours at 37° C. After 4 hours, ametabolic XTT assay was performed as outlined above.

Characterization of Bacterial Biofilm Adhesion on SLIPS-CoatedSubstrates. Freezer stocks of bacterial strains were maintained at −80°C. in 1:1 LB:glycerol. Overnight cultures of bacteria were grown inLuria-Bertani (LB) medium (P. aeruginosa and E. coli) or tryptic soybroth (S. aureus) at 37° C. with shaking at 200 rpm. Biofilms attachedto SLIPS-coated glass and bare glass substrates were imaged byfluorescence microscopy.

An inoculating subculture of P. aeruginosa was prepared bycentrifugation of the overnight culture at 4,000×g for 10 min followedby resuspension of the cell pellet in an amount of fresh M9+ medium,effecting a 1:10 dilution (v/v) of the overnight culture (M9+ mediumconsists of the M9 buffer, described above, supplemented with 0.4%arginine, 0.5% casamino acids, 0.2% glucose, 0.2% succinate, 0.2%citrate, 0.2% glutamate, 1 mM MgSO4, and 0.1 mM CaCl₂)) (see Frei etal., Angew. Chem. Int. Ed. 2012, 51, 5226). A subculture of S. aureuswas prepared by diluting overnight cultures 1:100 into fresh brain-heartinfusion medium supplemented with 1% (w/v) glucose (see Kratochvil etal., ACS Biomater. Sci. Eng. 2015, 1, 1039). E. coli subcultures wereprepared by diluting overnight cultures 1:1000 into fresh LB medium.

Both bare glass substrates and SLIPS-coated substrates were placedindividually into the wells of a 12-well microtiter plate (Costar 3737)and sterilized by UV irradiation for 20 minutes in a biological safetycabinet. Bacterial subculture (P. aeruginosa, S. aureus, E. coli) wasthen added to each well in 2 mL aliquots and the plates were incubatedunder static conditions at 37° C. for 24 h.

Substrates were then removed from the wells using forceps, gently dabbedon a paper towel to remove excess liquid, and placed in the wells of anew 12-well plate to perform biofilm staining with SYTO-9 according tothe manufacturer's protocol. Excess staining solution was removed bydabbing on a paper towel and the substrates were then transferred to thewells of a 24-well plate and covered by 400 μL PBS. Biofilms were thenimaged using an Olympus IX71 fluorescence microscope.

Characterization of Mammalian Cell Attachment on SLIPS-Coated Surfaces.All surfaces were sterilized prior to the seeding of cells by exposureto UV light for 15 minutes in a biological safety cabinet. Thesubstrates were then placed individually into the wells of 24-welltissue culture-treated polystyrene culture plates. HeLa cells werecultured in growth medium (MEM supplemented with 10% v/v fetal bovineserum, 100 units/mL penicillin and 100 μg/mL streptomycin), seeded onboth SLIPS-coated and bare glass substrates at initial densities of50,000 cells/mL in 750 μL of growth medium, and incubated at 37° C. for72 hours. Cytotoxicity measurements were conducted in replicates ofthree using a commercially available fluorescence live/dead assay kit(Molecular Probes).

For imaging, cells were stained with 500 μL of a staining solutioncontaining calcein AM and ethidium homodimer staining solution (1 μg/mLin PBS) for 45 minutes at 37° C. Following incubation, the stainingsolution was aspirated and replaced with 1 mL of fresh growth medium andcells were imaged by fluorescence microscopy.

Example 2—Prevention of Adhesion Using SLIPS Material

Multilayer-Based SLIPS Prevent Adhesion of C. albicans. It was recentlyreported that nanoporous and superhydrophobic polymer multilayersfabricated by the reactive layer-by-layer assembly of PEI and theamine-reactive polymer PVDMA can be infused with hydrophobic oils todesign surfaces that are ‘slippery’ to a range of aqueous fluids (seeManna et al., Adv. Mater. 2015, 27, 3007). The work described here (i)characterizes the ability of these slippery oil-infused surfaces toprevent the short- and longer-term attachment and colonization of fungaland bacterial cells, and (ii) tests the hypothesis that the porouspolymer matrix and liquid oil phases comprising these materials could beexploited as reservoirs for the loading and release of antimicrobialagents that could both improve the inherent antifouling properties ofthese surfaces and provide new strategies to kill non-adherent(planktonic) cells in surrounding media.

For all of the work described here, SLIPS fabricated by the infusion ofsilicone oil into nanoporous, decylamine-functionalized PEI/PVDMA films˜3.5 μm thick were used. Past studies by the present inventorsdemonstrate that SLIPS having this general structure and compositionexhibit water-droplet sliding angles as low as 1° and can be fabricatedon a broad range of objects, including the inner surfaces of tubes.

Silicone oil was used as a model hydrophobic oil, rather than the highlyfluorinated oils used by many other groups to design SLIPS in otherstudies (see Wong et al., Nature 2011, 477, 443; Epstein et al., Proc.Natl. Acad. Sci. U.S.A. 2012, 109, 13182; Leslie et al., Nat.Biotechnol. 2014, 32, 1134; and Xiao et al., ACS Appl. Mater. Inter.2013, 5, 10074), on the basis of the inventors' past results and becausesilicone oil can be used as a solvent for a range of differentsmall-molecule (drug-like) agents.

A series of initial experiments was performed to characterize theability of SLIPS-coated planar glass substrates to prevent fouling by C.albicans, an opportunistic human pathogen that is the leading cause ofhospital-acquired systemic fungal infections. These experiments wereperformed by incubating 50 μL droplets of C. albicans yeast inocula ontohorizontal bare (control) and SLIPS-coated glass surfaces for 3 hours at37° C. The droplets were then stained using crystal violet, which is adye used widely to stain microbial biomass and provide a visual means ofassessing the extent of macroscopic surface colonization (see Peeters etal., J. Microbiol. Meth. 2008, 72, 157; and Taff et al., Med. Mycology2012, 50, 214), and the substrates were tilted gradually from theirinitial horizontal positions to identify the angle at which the dropletsstarted to slide down the surfaces of the substrates (FIG. 1 (A)).

For SLIPS-coated glass substrates, a tilt angle of 10° was sufficient topromote sliding of beaded droplets of inocula, and sliding occurred downthe surface without leaving behind a trail of either liquid or yeastcells (FIG. 1 (B)). Furthermore, no staining was observed on thesurfaces of the SLIPS in the locations where the droplets were initiallyplaced. In contrast, droplets of yeast inocula spread readily on thesurfaces of bare glass substrates and required extreme tilt angles of90° to promote sliding (FIG. 1 (C)). On bare glass, sliding left behinda prominent trail of residual liquid, and upon washing the substrate weobserved adherent biomass to be present where the droplets of inoculawere initially placed.

To confirm these observations, a similar series of experiments wereperformed using the yeast-specific fluorescent dye FUN-1 to characterizethe presence or absence of live yeast cells. After tilting thesubstrates and washing them as described above, bright and uniform greenand red fluorescence were observed on the surfaces of bare glasssubstrates, indicating the presence of substantial numbers ofmetabolically active yeast cells (FIG. 1 (D-F); green is cytoplasmicstaining; red indicates intravacuolar structures in metabolically activecells). In contrast, the surfaces of SLIPS-coated glass slides were darkand almost completely devoid of fluorescence (FIG. 1 (G-I)).

When combined, these results demonstrate that glass surfaces coated withsilicone oil-infused porous PEI/PVDMA multilayers can substantiallyprevent the adhesion or attachment of C. albicans yeast for shortincubation periods.

Multilayer-Based SLIPS Resist the Formation of C. albicans Biofilms. Asecond series of experiments was performed to determine whether thesePEI/PVDMA-based SLIPS could prevent the attachment and colonization ofyeast for longer periods and thus prevent or reduce the formation of C.albicans biofilms.

For these experiments, SLIPS-coated glass substrates were immersed intothe wells of 24-well plates containing suspensions of C. albicans inmedia conditioned to stimulate biofilm formation, and incubated them at37° C. for 24 hours (bare glass substrates, bare glass smeared withsilicone oil (no polymer matrix), and multilayer-coated glass substrates(no oil) were also included as controls). After 24 hours, the substrateswere then either (i) stained directly with FUN-1 to visualize cellsattached to the surface or (ii) carefully removed and placed into emptywells to quantify the metabolic activity of attached cells using an XTTassay.

Robust biofilms exhibiting substantial metabolic activity were observedon all three control substrates (bare glass, oil-treated glass, andmultilayers without oil) by fluorescence microscopy (FIGS. 2 (A-C) and(G-L)) and the XTT assay (FIG. 2 (M)). In contrast, SLIPS-coatedsurfaces were almost completely devoid of biofilm (as determined byfluorescence microscopy; FIG. 2 (D-F) and exhibited very low metabolicactivity compared to the control surfaces (FIG. 2 (M)).

These results demonstrate that the individual components of these SLIPSthemselves (i.e., the porous polymer matrix and silicone oil) are notsubstantially cytotoxic, and suggest that the absence of biofilm onSLIPS-coated surfaces arises instead from slipperiness imparted by thefruitful combination of these two components. It is noted, however, thatwhile these materials can substantially prevent biofilm formation underthese conditions, small and isolated patches of biofilm wereoccasionally observed along the edges and corners of the coated glasssubstrates (FIG. 10) where defects may prevent uniform coverage by theSLIPS (these small patches of biofilm observed visually likely alsoaccount for the small amount of metabolic activity observed in our XTTassays; e.g., ˜6% as compared to biofilms formed on bare glasssubstrates; see FIG. 2 (M)). This observation is discussed further inthe sections below.

To characterize the anti-fouling properties of these SLIPS for longerperiods, a ‘multiple challenge’ experiment was performed in whichsubstrates were repeatedly (i) immersed in suspensions of yeast for 24hours, (ii) removed from the yeast suspensions and characterized, and(iii) immersed again in fresh yeast suspensions for another 24 hours(see the discussion above for additional details related to materialsand methods used in these experiments). This multiple challenge protocolwas adopted to circumvent limitations associated with the depletion ofnutrients (and resulting cell death) that would occur during prolonged,multiple-day immersion in a single yeast suspension. In addition to themultiple microbial challenges, this protocol also allowed potentialchanges in anti-fouling properties that could occur as a result of thephysical manipulation and re-use of SLIPS-coated surfaces to beassessed, including multiple passages through air/water interfaces.

After each 24-hour challenge, the substrates were removed tocharacterize slipperiness and extents of biofilm formation. FIG. 3 showsrepresentative results of an experiment involving three consecutivemicrobial challenges and reveals that SLIPS-coated surfaces maintainedtheir anti-fouling properties, compared to bare glass controls, asdetermined by bright-field and fluorescence microscopy characterizationof FUN-1-stained surfaces (FIG. 3 (A-H)). It is noted that the darkpunctate and granular structures observed in the bright-field images ofSLIPS-coated surfaces (FIG. 3 (B,D,F)) are associated with the texturesof the nanoporous multilayer scaffolds, and do not represent yeast cellsor biofilm (a result that is confirmed by the lack of green fluorescencein the FUN-1-stained SLIPS-coated surfaces shown in FIG. 3 (H)).

After the third microbial challenge, differences in the metabolicactivities of cells present on the surfaces of the substrates (FIG. 3(I)) were also quantified. SLIPS-coated substrates exhibited an ˜80%reduction in biofilm formation as compared to bare glass substrates.Based on this result and visual inspection of FUN-1-stained samples, the˜20% metabolic activity observed here (relative to controls) wasattributed to the presence of small patches of biofilm located at thecorners and edges of the SLIPS-coated substrates, as described above,and not to the presence of biofilm over the remaining majority of theliquid-infused surfaces.

Characterization of Anti-Fouling Behavior in SLIPS-Coated Catheter TubeSegments. One useful feature of the layer-by-layer process used tofabricate the materials reported here is that it permits the fabricationof SLIPS on a variety of types of surfaces, including plastic and metalsurfaces and flexible, curved, or topologically complex surfaces (e.g.,tubing) typical of devices or equipment relevant in applied industrialor biomedical contexts (see Huang et al., ACS Macro Lett. 2013, 2, 826;Manna et al., Adv. Mater. 2015, 27, 3007; and Sunny et al., Adv. Funct.Mater. 2014, 24, 6658). The results of experiments shown in FIG. 11demonstrate that multilayer-based SLIPS can also prevent C. albicansbiofilm formation on the surfaces of flexible plastic (PET; FIG. 11(E-H)) and metal (aluminum foil; FIG. 11 (I-L)) surfaces.

A series of experiments was also performed to characterize the abilityof SLIPS to prevent C. albicans adhesion and biofilm formation on theinner surfaces of narrow diameter polymer-based catheter tubes. FIG. 4(A-B) shows low- and high-magnification SEM images of micro/nanoporousPEI/PVDMA multilayers on the inner surfaces of polyethylene tubes ˜850μm in diameter (images were acquired prior to the infusion of siliconeoil). The images in panels C-E of FIG. 4 demonstrate that the innersurfaces of these tubes become slippery when infused with oil. Theseimages show a 5 μL aliquot of an aqueous solution of the red dyetetramethylrhodamine (TMR) slipping rapidly through a SLIPS-coatedcatheter tube held at a 10° angle (FIG. 4 (F)) shows an image of asimilar experiment conducted using a bare, uncoated tube; the liquid inthis case remains pinned at the top of the tube and does not passthrough the tube under these conditions).

Segments of uncoated and SLIPS-coated catheter tubes were filled with C.albicans inocula, incubated at 37° C. for 4 hours, and then used SEM tovisualize and characterize the amounts and types of cells adhered toboth types of tubes. Hyphal and pseudohyphal C. albicans cells wereobserved on the surfaces of bare control tubes (FIG. 4 (I-K)), asexpected in the early stages of biofilm formation. The surfaces ofSLIPS-coated catheters were largely devoid of C. albicans cells (FIG. 4(L-N)); the roughness and texture apparent in panels M and N are againfeatures associated with the polymer multilayer matrix; e.g., see FIG. 4(A); FIG. 12 shows representative fluorescence microscopy images ofFUN-1-stained bare or SLIPS-coated samples).

The metabolic activity of cells associated with SLIPS-coated surfaceswas also ˜85% lower than that measured in bare, uncoated catheter tubes(as determined by an XTT assay; FIG. 4 (G)). Furthermore, it wasestimated that ˜5 times more planktonic cells remained in the liquidmedia contained inside SLIPS-coated tubes in these experiments (FIG. 4(H)) compared to the media inside bare, uncoated tubes that permit orpromote cell attachment. This result is to be expected in view of theanti-fouling nature of the SLIPS-coated tubes (to which suspended,planktonic cells cannot attach). This result also illustrates, however,one potential limitation of SLIPS in certain contexts—namely that SLIPScan prevent or reduce microbial cell attachment and biofilm formation onthe surfaces to which they are applied, but can themselves do little toprevent the growth and proliferation of planktonic C. albicans cells andother potentially harmful pathogens in surrounding aqueous environments.

The results above also demonstrate that the anti-fouling nature of theseSLIPS is not perfect and, in particular, that fungal adhesion andbiofilm formation can occur over short periods in locations (e.g., edgesand corners) where slippery character may be compromised. It isconsidered likely that this latter issue could be addressed through theapplication of SLIPS to substrates that do not contain sharp edges, suchas the glass slides used in several of the above experiments. In thesections below, however, a controlled release approach is described fornew multi-functional SLIPS that could help address these broader issuesand expand the potential utility of these slippery surfaces.

Multilayer-Based SLIPS Prevent Adhesion & Colonization by Bacteria &Mammalian Cells. The ability of silicone oil-infused, PEI/PVDMA-basedSLIPS-coated surfaces to resist attachment and fouling by commonbacterial pathogens and mammalian cells was also investigated. For theseexperiments, SLIPS-coated glass substrates and bare glass slides insuspensions of one Gram-negative (P. aeruginosa) and two Gram-positive(E. coli and S. aureus) species of bacteria were incubated for 24 hoursat 37° C. All surfaces were then removed from solution and stained usinga SYTO-9 biofilm staining solution to identify the presence of bacterialbiofilms using fluorescence microscopy.

The images in FIG. 5 (A-F) clearly indicate the presence of adherent E.coli and dense biofilms of P. aeruginosa and S. aureus on bare glasssubstrates (FIG. 5 (A,C,E)). These images also reveal the lack ofbacterial colonization and biofilm formation on SLIPS-coated substrates(FIG. 5 (B,D,F)). Finally, FIG. 5 (G-H) show images of SLIPS-coated andbare glass surfaces after incubation with HeLa cells, a human cervicalcarcinoma cell line widely used as a model in biomedical research, at37° C. for 72 hours (cells were stained using a calcein AM fluorescentcell stain prior to imaging).

The fluorescence microscopy images in FIG. 5 (G-H) demonstrate thatSLIPS coatings can also substantially prevent attachment and fouling bythis mammalian cell line. FIG. 13 shows additional representative imagesassociated with these experiments. Overall, these results demonstratethat these multilayer-based SLIPS coatings can prevent surface foulingby fungal, bacterial, and mammalian cells under a range of differentconditions and time scales.

Example 3—Effects of SLIPS Material on Planktonic Cells

SLIPS Loaded with Antimicrobial Agents Prevent Fouling and KillPlanktonic Cells. One guiding hypothesis of the work described herein isthat the infused oil phases of these multilayer-based SLIPS can be usedto host and sustain the release of small-molecule antimicrobial agents.It is asserted that if antimicrobial agents are incorporated withoutdegrading the inherent slippery character of these surfaces, the releaseof these agents into surrounding liquid media could kill planktoniccells and further prevent or reduce the likelihood of biofilm growth.

To explore the feasibility of this approach and establishproof-of-concept, experiments were performed using silicone oil-infusedSLIPS and triclosan, a model broad-spectrum antimicrobial agent that cankill both fungal and bacterial cells (the molecular structure oftriclosan is shown in the inset of FIG. 6 (C)). Triclosan is soluble insilicone oil and thus permits the facile design of triclosan-loadedSLIPS by direct infusion of triclosan/silicone oil solutions intonanoporous multilayer matrices. In this study, however, an alternativeapproach to loading was adopted that involves the solvent-assistedloading of triclosan into the nanoporous multilayers prior to infusionwith silicone oil (see discussion above for additional details relatedto materials and methods used in these experiments).

This technically straightforward, solvent-assisted method was used forthree reasons: (i) it allows precise control over the amount oftriclosan (or any other anti-microbial agent) loaded, (ii) it permitsthe loading of agents into SLIPS at concentrations that far exceed theirsolubility in the oil phase and, perhaps most notably, (iii) it has thepotential to promote more gradual and sustained release. Without beingbound by theory, it is believed the sustained release involves thepartitioning of triclosan from the polymer matrix into the oil and fromthe oil into the surrounding aqueous phase, which provided a moresustained release than a material in which all of the loaded agent isinitially present in the oil phase.

FIG. 6 shows the results of experiments using triclosan-loaded SLIPSprepared by the pre-oil-infusion treatment of porous PE I/PVDMAmultilayers with solutions of triclosan in acetone. As shown in FIG. 6(A), the loading of triclosan into the polymer matrix did not have asubstantial impact on the advancing water contact angle (Bath) forloadings of triclosan up to 250 μg/cm² (θadv ˜˜103°), and reduced itonly slightly at loadings of 500 μg/cm² (θ_(adv) ˜93°). The contactangle hysteresis (θ_(hys)) of these surfaces increased gradually over asmall range (from ˜3° to ˜8°) as triclosan loading was increased from 0μg/cm² to 500 μg/cm² (FIG. 6 (A)). This combination of features allowstriclosan to be incorporated into oil-infused SLIPS that maintain theirslippery properties, as demonstrated by sliding of a 15 μL droplet of anaqueous TMR solution on the triclosan-loaded SLIPS-coated glass slideshown in FIG. 6 (B) (tilt angle=10°; loading=500 μg/cm²).

FIG. 6 (C) shows the cumulative amount of triclosan released intosolution over time for triclosan-loaded SLIPS submerged and incubated inPBS buffer at 37° C. (closed circles; open circles correspond to therelease profile of otherwise identical films not loaded with triclosan).Approximately 30% of the loaded triclosan was released over the first 20days of incubation, with an additional ˜15% of the triclosan releasedover an additional 100 days. This profile is consistent with an initialburst release phase followed by a slower and more sustained phase ofrelease, and is also consistent with concentration dependent diffusionprocesses involving the gradual partitioning of triclosan from thepolymer matrix into the oil phase and into solution.

These results demonstrate that this loading approach can be used todesign SLIPS that promote the sustained release of an antimicrobialagent for a period of at least 4 months. It is noted that only ˜45% ofthe initially loaded triclosan was released over this 4-month period.The results shown in FIG. 6 (C) suggest that release would likelycontinue occur over a substantially longer period, but the release wasnot monitored for longer than 4 months as part of this proof-of-conceptstudy.

To determine whether the triclosan released from these SLIPS could bereleased in amounts sufficient to kill planktonic C. albicans andinhibit biofilm formation, triclosan-loaded SLIPS-coated glass slideswere immersed in C. albicans inocula and incubated at 37° C. for 24hours (SLIPS-coated glass substrates without triclosan and bare glasssubstrates were used as controls). The amounts of metabolically activeyeast present both (i) on the surfaces of the substrates and (ii) in thesurrounding media in the wells containing the substrates were quantifiedseparately using the XTT assay; the quantitative results of thesestudies are shown in FIG. 7 (representative qualitative visual resultsare also shown in FIG. 14).

Triclosan-loaded SLIPS exhibited substantial and significant reductionsin the amounts of C. albicans present both in solution and on theirsurfaces relative to SLIPS without triclosan and bare glass substrates.In particular, the nearly complete lack of metabolically active C.albicans present in solution when triclosan-loaded SLIPS were used (FIG.7; right) demonstrates that the release of triclosan occurred in amountssufficient to kill nearly all of the planktonic C. albicans cellspresent in the liquid media.

The loading of triclosan also yielded significant reductions in theamount of metabolically active yeast associated with the surfaces of theSLIPS (FIG. 7; left), a result that is likely a direct outcome of theability of those SLIPS to substantially reduce the number of viableplanktonic C. albicans cells in their surrounding environments. Becausetriclosan-loaded SLIPS release triclosan for prolonged periods, amultiple challenge experiment was performed similar to that describedabove as well as a second, longer-term incubation experiment todetermine whether triclosan-loaded SLIPS could exhibit enhancedresistance to C. albicans colonization under longer and more challengingconditions.

For multiple challenge experiments, triclosan-loaded SLIPS (and controlbare glass substrates) were subjected to five sequential 24-hourimmersions in wells containing freshly prepared C. albicans inocula. Asshown in FIG. 8 (A), SLIPS loaded with triclosan were observed toprevent biofilm formation on these surfaces (it was also observed thatalmost no viable cells were in solution at the remainder of each 24-hourchallenge, consistent with the results shown in FIG. 7; data formultiple challenges not shown here).

For the long-term antifungal experiment, substrates were placed in wellscontaining C. albicans inoculum and removed at the end of one week (withone quarter of the volume of the culture media in the wells gentlyreplaced every two days). Triclosan-loaded SLIPS were observed to havesubstantially reduced amounts of metabolically active C. albicans cellsassociated with their surfaces after this extended period of incubation(compared to glass controls; FIG. 8 (B)).

In a final series of experiments, triclosan-loaded SLIPS were alsofabricated inside catheter tubes. As shown in FIG. 9, the incorporationof triclosan also resulted in substantial reductions in both biofilmsand viable cells in the intraluminal solutions of media incubated inthese tubes.

When combined, the results of these experiments demonstrate that thesemultifunctional, triclosan-loaded SLIPS can exert influences inimportant ways that extend beyond those that rely on direct interactionswith cells at their slippery surfaces. These results also demonstratethat this approach to killing planktonic microbial cells in surroundingmedia can improve the anti-fouling properties of these materials.Because triclosan is a broad-spectrum antimicrobial agent that alsoexhibits substantial activity against bacteria, it is anticipated thatthe approach to the loading and release of triclosan described above forthe design of SLIPS that kill planktonic fungal cells could also be usedto improve the antibacterial properties of these coatings. This approachthus has the potential to be general and provide broader benefits inbiomedical contexts by eliminating both planktonic yeast and bacterialcells that could promote downstream infections or produce harmful toxins(e.g., hemolysin, toxic shock syndrome toxin, etc.) through processesthat are independent of problems associated with the simple attachmentof cells or the formation of biofilms on surfaces.

Summary and Conclusions. The above examples demonstrate an approach tothe design of slippery liquid-infused porous surfaces (SLIPS) that canboth strongly prevent surface fouling and effectively kill microbialpathogens in surrounding media. This approach addresses a currentlimitation of SLIPS-based coatings reported by other groups, which canprevent fouling by microorganisms on the surfaces to which they areapplied, but cannot prevent the proliferation of or kill nearbynon-adherent cells that could colonize nearby surfaces or engage inother behaviors that could lead to infections or other associatedburdens. The multi-functional SLIPS reported here address theseimportant issues and thus have the potential to significantly expand theutility and effectiveness of these anti-fouling surfaces in a range ofbiomedical, industrial, and commercial contexts.

These results demonstrate that SLIPS fabricated by the infusion ofsilicone oil into nanoporous polymer multilayers can prevent short- andlonger-term colonization and the formation of biofilms by the prevalentand opportunistic fungal pathogen C. albicans. These results alsodemonstrate that the porous polymer and hydrophobic oil phasescomprising these materials can be exploited to load and then sustain therelease of the hydrophobic and broad-spectrum antimicrobial agenttriclosan into surrounding media. Using the silicone oil-infusedPEI/PVDMA model SLIPS system reported here, these results demonstratethat the release of triclosan into surrounding aqueous media can besustained for extended periods (up to at least 4 months, the longestperiod investigated in this proof-of-concept study). This approachimproves both the inherent anti-fouling properties of these slipperysurfaces (e.g., by reducing surface-associated fungi and biofilm growththat can occur at defects present at the edges of planar substrates)and, importantly, endows these coatings with the ability to efficientlykill planktonic C. albicans.

Finally, these results demonstrate that these SLIPS coatings can alsoprevent surface fouling by common bacterial pathogens and a modelmammalian cell line and that they can be fabricated on the insides offlexible catheter tubes, suggesting one potential applied biomedicalcontext in which the multi-functionality of this dual-action approachmay prove useful. It should be noted, however, that the general strategyreported here for the loading and release of small-molecule agents intoslippery surfaces is not likely to be limited to the model systemreported here, and has the potential to be general. Provided that theproperties of the porous matrix and the infused oil are chosenappropriately, for example, this general approach could be used to hostand control the rates of release of a broad range of functional agentsinto aqueous environments or other surrounding media. As such, thematerials, strategies, and concepts reported here have the potential toopen the door to many new applications of this new class of slipperyliquid-infused materials.

Example 4—Fabrication of SLIPS Material with Quorum Sensing Modulators

Embodiments of the present invention seek to further develop thepotential of SLIPS as reservoirs for the controlled release of activeagents, with a focus on the design of multifunctional andchemical-eluting SLIPS capable of attenuating the colonization andvirulence of planktonic bacteria through non-biocidal means (e.g., bysustaining the release of active agents that do not necessarily killbacteria, but instead attenuate virulent behaviors by targetingnon-essential pathways). Such ‘anti-virulence’ strategies have attractedconsiderable interest over the past decade as the incidence of bacterialresistance has increased (see Clatworthy et al., Nat. Chem. Biol. 2007,3, 541-548; and Allen et al. Nat. Rev. Microbiol. 2014, 12, 300-308).

One promising target for potential anti-virulence approaches isbacterial quorum sensing (QS) circuits. QS is a small molecule-basedcommunication system used by many bacteria to coordinate the expressionof group-beneficial behaviors when a threshold population density (i.e.,a ‘quorum’) is reached (see Bassler et al., Cell 2006, 125, 237-246;Camilli et al. Science 2006, 311, 1113-1116; and Ng et al., Annu. Rev.Genet. 2009, 43, 197-222). In many common pathogens, such as theGram-negative bacterium Pseudomonas aeruginosa, QS systems control theproduction of excreted virulence factors and the formation of biofilms,but are non-essential for cell growth—targeting these systems thuspresents a basis for the development of ‘non-biocidal’ approaches tocontrolling bacterial virulence. Over the last 10 years, many potentsmall molecule inhibitors of QS (QSIs) that are active in P. aeruginosaand other pathogens, and that represent valuable chemical tools to testsuch anti-virulence approaches, have been developed (Eibergen et al.,Chem BioChem 2015, 16, 2348-2356; Geske et al., Bioorg. Med. Chem. Lett.2008, 18, 5978-5981; Geske et al. ChemBioChem 2008, 9, 389-400; Mattmannet al. ChemBioChem 2011, 12, 942-949; and Stacy et al., ACS Chem. Biol.2012, 7, 1719-1728).

As part of a broader effort to develop and exploit the therapeuticpotential of QSIs, strategies have been developed for the encapsulationor integration of QSIs and other anti-virulence agents intopolymer-based materials or onto inorganic surfaces (Breitbach et al.,Chem. Comm. 2011, 47, 370-372; Broderick et al., 2013, Adv. HealthcareMater. 2, 993-1000; Broderick et al., Adv. Healthcare Mater. 2014, 3,97-105; and Manna et al., 2013, Adv. Mater. 25, 6405-6409; Kratochvil etal., ACS Biomater. Sci. Eng. 2015, 1, 1039-1049.).

These past studies have yielded many different approaches to the releaseof anti-virulence agents, but they have relied, in large measure, onmaterials and tactics that do not inherently prevent biofouling (apartfrom the activities of the released inhibitors). The present exampledemonstrates that the polymer and oil phases of polymer-based SLIPS canbe exploited to load and control the release of synthetic smallmolecules that inhibit or modulate QS in P. aeruginosa. It is alsodemonstrated that these QSIs can be loaded into SLIPS without affectingslippery or anti-fouling properties, and that the agents remainbiologically active, enabling QSI-loaded SLIPS to both prevent bacterialcolonization and attenuate important QS-regulated behaviors, such as theproduction of key excreted virulence factors, in planktonic cultures ofthis pathogen. These liquid-infused materials can also be designed torelease multiple QSIs that target multiple different QS circuitssimultaneously.

Finally, it is demonstrated that these polymer-based SLIPS, which areinherently resistant to the formation of P. aeruginosa biofilms on theirown surfaces, can also release non-bactericidal biofilm inhibitors thatconfer robust protection against the formation of biofilms on othernearby and unprotected (non-SLIPS-coated) surfaces. These resultssuggest the basis of new non-bactericidal approaches to the design andprotection of anti-fouling surfaces that circumvent critical problemsassociated with the use of antibiotics. More broadly, this work alsoadvances new approaches to the integration of controlled releasestrategies with SLIPS-based technologies that could improve theproperties of these inherently anti-fouling, oil-infused surfaces in arange of other contexts.

Materials and General Considerations. All chemicals were purchased fromSigma-Aldrich, unless indicated otherwise, and used without furtherpurification. 2-Vinyl-4,4-dimethylazlactone (VDMA) was a gift from 3MCorporation, Minneapolis, Minn. Poly(2-vinyl-4,4-dimethylazlactone)(PVDMA) was synthesized as described previously (Buck et al., Chem.Mater. 2010, 22, 6319-6327). Glass microscope slides were purchased fromFisher Scientific (Pittsburgh, Pa.). Dimethyl-2-aminobenzamidazole(DMABI), and compounds C₁₄, E22, and V-06-018 were synthesized aspreviously reported (Frei et al., Angew. Chem. Int. Ed. 2012, 51,5226-5229; Geske et al., ChemBioChem 2008, 9, 389-400; Geske et al.,Bioorg. Med. Chem. Lett. 2008, 18, 5978-5981; and Muh et al.,Antimicrob. Agents Chemother. 2006, 50, 3674-3679). Compressed air usedto dry samples was filtered through a 0.2 μm membrane syringe filter.UV/vis measurements were made using a Beckman Coulter DU520 UV/visspectrophotometer (Fullerton, Calif.). Fluorescence microscopy imageswere acquired using an Olympus IX70 microscope and analyzed using theMetavue version V7.7.8.0 software package (Molecular Devices). Log Pvalues were calculated using Perkin Elmer ChemDraw version 13.0.0.118(Cambridge Soft Corporation). Absorbance measurements in biologicalassays were made using a Biotek Synergy 2 plate reader running Gen 5software (version 1.05).

Fabrication of Polymer Multilayers. Prior to film fabrication, glassmicroscope slides were cut into 0.8 cm wide strips and scored in 1 cmlong segments. Covalently crosslinked and nanoporous polymer multilayerscomposed of PVDMA and branched poly(ethyleneimine) (BPEI; MW ˜25,000),referred to from hereon as BPEI/PVDMA multilayers, were fabricated onthe glass substrates using a covalent/reactive layer-by-layer assemblyprocess, as previously described (Manna et al., Adv. Mater. 2015, 27,3007-3012). Briefly, the substrates were submerged iteratively in thefollowing solutions for 20 seconds each: (i) BPEI (20 mM in acetone withrespect to the polymer repeat unit); (ii) two acetone rinses; (iii)PVDMA (20 mM in acetone with respect to the polymer repeat unit); (iv)two additional acetone rinses. This cycle was repeated 35 times. Polymersolution volumes were maintained with fresh acetone to compensate forevaporation and maintain polymer concentration during the dippingprocess. After fabrication, films were functionalized and renderedhydrophobic by immersion in a 20 mM solution of n-decylamine in THFovernight at room temperature. Functionalized films were then rinsed anddried using compressed, filtered air. Film-coated substrates were thenfragmented along the pre-fabrication scores to produce 0.8 cm×1.0 cmsamples.

Small Molecule Loading, Oil Infusion, and Characterization of Release. A10 μL aliquot of an acetone solution of a small-molecule agent (12.0 mMin compounds E22, C₁₄, or V-06-018 for individual release experiments;6.0 mM of both compound E22 and compound V-06-018 for dual releaseexperiments; 15.0 mM for experiments using DMABI) was applied to the topside of a film-coated substrate, allowed to dry, and repeated on thecoating on the opposite side of the substrate. Immediately prior to use,loaded films were infused with silicone oil (for melting point andboiling point apparatuses; Sigma-Aldrich) by placing a 2.25 μL dropletof oil on each side and allowing the oil to spread over the entiresurface. Excess oil was removed from the surface using tissue paper. Forrelease experiments, small-molecule-loaded and oil-infused films wereincubated at 37° C. in 1.0 mL of PBS buffer (pH=7.4). At designated timepoints, substrates were removed from the incubation buffer and placed infresh buffer before returning to the incubator. The release of theloaded agents into the incubation buffer was characterized using aUV/vis spectrophotometer.

Bacterial Strains and Growth Conditions. All media and reagents forbacterial culture were purchased from commercial sources. Wild-type P.aeruginosa strain PAO1 was obtained from the University of Rochester.Overnight cultures of bacteria were grown in Luria-Bertani (LB) mediumat 37° C. with shaking at 200 rpm. Freezer stocks of bacterial strainswere maintained at −80° C. in 1:1 LB:glycerol. MOPS glutamate wasprepared as described by Mellbye et al., J. Bacteriol. 2014, 196,1155-1164. The assay medium was prepared prior to each experiment bydiluting 10×MOPS buffer (500 mM MOPS, 40 mM tricine, 500 mM NaCl, 10 mMK₂HSO₄, 500 μM MgCl₂, 100 μM CaCl₂), 3 μM (NH₄)₆Mo₇O₂₄, 400 μM H₃BO₃, 30μM Co(OAc)₂, 10 μM CuSO₄, 80 μM MnSO₄, 10 μM ZnSO₄, pH 7.0, filtersterilized) into sterile 18 MΩ water. To this working solution, asterile 10× stock solution of L-glutamate (250 mM) and sterile 100×stock solutions of K₂HPO4 (400 mM), FeSO₄ (500 μM), and NH₄Cl (1.5 M)were added in appropriate amounts.

Pyocyanin Assay Protocol. The amount of pyocyanin in P. aeruginosaculture supernatants was measured following the protocol of O'Loughlinet al. Proc. Natl. Acad. Sci. U.S.A 2013, 110, 17981-17986, withmodifications. A 10 mL overnight culture of P. aeruginosa PAO1 was grownfor 16 h. An inoculating culture was prepared by diluting the overnightculture 1:100 into freshly prepared MOPS glutamate medium, and 2 mLaliquots of this subculture were added to each test tube (0.5% DMSO).SLIPS-coated surfaces (sterilized by UV irradiation for 20 min in abiological safety cabinet) were placed in each tube, and the cultureswere grown for 17 h at 37° C. with shaking incubation at 200 rpm. Thefinal cell density was measured by reading absorbance at 600 nm (OD600).Relative pyocyanin levels were measured by first pelleting 1.5 mL ofwell-mixed culture at 4,000×g for 10 min, transferring 200 μL of theresulting supernatant to a clear, plastic 96-well microtiter plate, andreading absorbance at 695 nm. Media background absorbance (measured froma “no bacteria” control) was subtracted, the resulting values werenormalized by dividing by the final OD600, and the data were plottedrelative to an unloaded positive SLIPS control.

P. aeruginosa Biofilm Growth and Crystal Violet Staining Assay Protocol.Biofilm formation by P. aeruginosa was quantified by crystal violet (CV)staining following the protocol of Frei et al., Angew. Chem. Int. Ed.2012, 51, 5226-5229 with modifications. A 10 mL overnight culture of P.aeruginosa PAO1 was grown for 16 h. An inoculating subculture wasprepared by centrifugation of the overnight culture at 4,000 g for 10min followed by resuspension of the cell pellet in an amount of freshM9+ medium (see Frei et al. for full details of this medium)supplemented with 5% (v/v) LB to effect a 1:10 dilution (v/v) of theovernight culture.

Glass substrates were placed into the wells of a 12-well microtiterplate (Costar 3737) and sterilized by UV irradiation for 20 min in abiological safety cabinet. Subculture was added to each well in 2 mLaliquots and the plates were incubated under static conditions at 37° C.for 24 h. Substrates were removed from the wells using forceps, gentlydabbed on a paper towel to remove excess liquid, and placed in a new12-well plate. Spent culture medium was removed from the wells byinverting the assay plate over a basin and the attached biofilm waswashed once with 1 mL of PBS. The substrates and assay plate wereallowed to dry in a 37° C. incubator for at least 8 h. The substratesand well bottoms were stained with 1 mL of a CV solution (0.1% CV (w/v)in 95:5 water:ethanol) for 10 min. Excess CV stain was removed bywashing twice with 1 mL of water, and the substrates and plate weredried at 37° C. for at least 4 h. CV stain absorbed by the attachedbiofilm was quantified by re-solubilizing the stain in 1 mL (wells) or0.5 mL (substrates) 30% (v/v) acetic acid, transferring 200 μL of thissolution to a clear 96-well microtiter plate (Costar 3370), andmeasuring absorbance at 590 nm.

Characterization of Biofilms Using Fluorescence Microscopy. Biofilmsattached to glass substrates were imaged by fluorescence microscopyusing the above biofilm growth protocol with the followingmodifications. After incubation, substrates were gently removed from theassay medium using forceps, washed once by dipping into PBS, and stainedwith SYTO 9 (Invitrogen) according to the manufacturer's protocol.Excess staining solution was removed by dabbing on a paper towel and thesubstrates were covered by 400 μL of PBS in a 24-well plate. Biofilmswere then imaged using an Olympus IX71 fluorescence microscope.

Fabrication and loading of QSIs into nanoporous, multilayer-based SLIPS.The SLIPS used in this study were fabricated by the infusion of siliconeoil into nanoporous and covalently-crosslinked polymer multilayersfabricated by the reactive layer-by-layer assembly ofpoly(vinyl-4,4-dimethylazlactone) (PVDMA) and branchedpoly(ethyleneimine (BPEI) on planar glass substrates. After filmfabrication, these reactive multilayers were treated with n-decylamineto functionalize residual azlactones remaining in the films withhydrophobic alkyl groups (FIG. 15 (A)) and render them more chemicallycompatible with silicone oil. Infusion of silicone oil into thedecylamine-functionalized multilayers yielded SLIPS that exhibited waterdroplet sliding angles of ≤10° in agreement with past studies.

Two P. aeruginosa QSIs [E22 (an acyl L-homoserine lactone (AHL)-basedantagonist of the RhIR QS receptor) and V-06-018 (a non-AHL-basedantagonist of the LasR QS receptor)] and DMABI (a potent biofilminhibitor that modulates QS in P. aeruginosa through a yet to bedetermined mechanism) were selected for this study (FIG. 16) becausethey represent some of the most potent QS modulators known.

QSI- and DMABI-loaded SLIPS were prepared by applying a 10 μL droplet ofa 12 mM acetone solution of QSI or a 15 mM acetone solution of DMABI toeach side of the multilayer-coated substrates (prior to the infusion ofoil; FIG. 15 (B)). The acetone solutions quickly wet the entirety of thenanoporous coatings and, upon evaporation, left the loaded compoundsadsorbed within the multilayers. A small excess of silicone oil was thenpipetted onto both sides of the compound-loaded coatings and allowed toinfuse and spread across the entirety of the surface (FIG. 15 (C)). Thisapproach to loading was adopted (e.g., as opposed to an alternativeapproach in which dry multilayers were infused directly with siliconeoil containing dissolved compound) on the basis of our past resultsusing triclosan, and because this approach enables more precise controlover the amount of compound loaded in ways that are not restricted bythe solubility of a given compound in the oil phase.

All QSI-loaded SLIPS used in this study contained 240 nmol of compoundper substrate and all DMABI-loaded SLIPS contained 300 nmol of compound.It was confirmed that the loading of these small molecules did notimpact the slippery properties of the resulting oil-infused multilayersby placing 10 μL droplets of water on QSI-loaded SLIPS held at a tiltangle of 10° and measuring the sliding velocities of the droplets. Asrevealed by the results shown in Table 1, SLIPS loaded with E22,V-06-018, DMABI, or a 1:1 ratio of both E22 and V-06-018 (at a loadingof 120 nmol each) did not have a substantial impact on droplet slidingvelocities.

TABLE 1 Impact of Loading on Sliding Velocities Loaded Compound SlideVelocity (mm/s)^(a) None 7.6 ± 0.3 C14 8.4 ± 0.6 E22 6.9 ± 0.3 V-06-0186.6 ± 0.6 E22 + V-06-018 4.3 ± 0.2 DMABI 7.4 ± 0.2 ^(a)Values are themean and standard error of six independent replicates; experiments wereperformed by placing droplets of water on SLIPS-coated surfaces held atangles of 10° and measuring the times required to slide a fixed distance(see Materials and Methods for details).

Characterization of the release of QSIs and biofilm inhibitors fromSLIPS. QSI-loaded SLIPS were incubated in phosphate-buffered saline(PBS) to characterize the release of the imbedded QSIs into surroundingmedia under physiologically relevant conditions (37° C.; pH=7.4). Forthese studies, SLIPS loaded with compound E22 and DMABI were usedbecause the relatively strong UV absorbance of both compounds permittedfacile monitoring of release by UV/vis spectrophotometry (the absorbanceof compound V-06-018 was significantly obscured by the presence ofsilicone oil in these experiments; release of this compound was thus notevaluated quantitatively).

As shown in FIG. 17, compound E22 was released into surrounding bufferrelatively quickly, with approximately 75% of the total amount loadedreleased over the first 12 hours of incubation. No further compound wasreleased over an additional 190 hours of incubation, suggesting that˜25% of this compound remained strongly bound to the polymer matrix.Additional experiments using otherwise identical SLIPS loaded with AHLderivative C14 (structure shown in FIG. 16) revealed this more polar AHLto be released almost completely over a period of 24 hours. Finally, incontrast to the relatively rapid release exhibited by these AHL-loadedSLIPS, coatings loaded with the 2-aminobenzimidazole-based biofilminhibitor DMABI released their contents much more slowly, withapproximately 40% of the loaded compound released after the first 24hours, and an additional ˜40% released over the next 150 hours; see FIG.17).

It is clear from these results that the structure of the loadedcompounds can have a significant influence on rates and extents ofrelease, likely a result of differences in the interactions of thesecompounds with the porous polymer matrix and differences in the extentto which the compounds partition into the silicone oil phase. Additionalexperiments are underway to better understand the factors that lead tothese large differences, as well as the extent to which changes in filmstructure and the properties of infused oil phase can be exploited totune the release profiles of these and other agents more broadly. Thetime scales and the amounts of QSIs and DMABI released by the materialsreported here (FIG. 17) were more than sufficient to demonstrate robustproofs of concept in all biological studies described below.

Release of QSIs from SLIPS attenuates P. aeruginosa pyocyaninproduction. To characterize the biological activities of QSI-loadedSLIPS, production of the redox-active virulence factor pyocyanin in P.aeruginosa was monitored. Pyocyanin production is controlled by the QSreceptors LasR and RhIR in P. aeruginosa, and should thus be attenuatedby both V-06-018 and E22. Wild-type P. aeruginosa was grown in thepresence of SLIPS substrates loaded with the LasR antagonist V-06-018 orthe RhIR antagonist E22 (in amounts designed to yield approximately 100μM of compound in the assay culture upon full release) and quantifiedpyocyanin production after 17 hours of shaking incubation.

As shown in FIG. 18, SLIPS loaded with V-06-018 and E22 inhibitedpyocyanin production by approximately 80% and 45%, respectively. Thesevalues are equivalent to the levels of pyocyanin inhibition observedwhen these compounds are administered exogenously, indicating that bothof these compounds are released from the SLIPS-coated surfaces inintact, biologically-active forms and in concentrations sufficient toinhibit QS under these assay conditions. Although not investigatedspecifically as part of this study, it is noted that the overallstrategy used here, in which encapsulated payloads are stored within apolymer matrix infused with a hydrophobic and water-immiscible oil—and,thus, largely protected from contact with bulk water until they diffuseacross the oil/water interface—could prove useful for the prolongedrelease of active agents that hydrolyze or decompose readily uponcontact with water. For instance, it is well known that AHLs hydrolyzerelatively rapidly in aqueous media and that the ring-opened forms areinactive; we anticipate that SLIPs containing AHLs (such as E22 and C12)could provide means for extending their effective half-lives in water,and thus extending their utility as QSIs.

It was previously demonstrated that cocktails of compounds targetingmultiple different QS circuits could result in greater attenuation ofvirulence factor production (as compared to levels attenuated upon theadministration of a single compound targeting a single QS circuit) (seeWelsh, M. A., and Blackwell, H. E. (2016) Chemical genetics revealsenvironment-specific roles for quorum sensing circuits in Pseudomonasaeruginosa, Cell Chem. Biol. 23, 361-369). To explore the potential ofour polymer-based SLIPS to promote the simultaneous release of twodifferent active agents, and develop SLIPS-coated surfaces thatattenuate QS more strongly than those described above, SLIPS wereprepared loaded with both V-06-018 and E22 (in amounts designed to giveapproximately 50 μM of each compound in the assay culture upon fullrelease) and pyocyanin production was quantified when the compounds werereleased simultaneously. Over 90% inhibition of pyocyanin production wasobserved using this dual-release approach (FIG. 18). This dual-QSIrelease approach allows for lower loadings of each individual agent andpromotes levels of inhibition greater than those exhibited when eitheragent is used alone because it targets both RhIR and LasR QS receptorssimultaneously.

The ease with which these QSIs (and other acetone-soluble agents) can beloaded into these nanoporous multilayers, without necessitating anychanges to the fabrication process, suggests that this basic approachshould be general and that these SLIPS systems should be appropriate foruse in other applications that would benefit from the simultaneousrelease of single or multiple different active agents.

It is noted that the SLIPS-coated substrates emerging from these in situvirulence factor production experiments remained anti-fouling tobacteria, but exhibited water droplet sliding angles higher than thosethat were measured prior to incubation with bacteria (e.g., droplets ofwater required tilt angles of ˜30° or more to slide freely, as comparedto sliding angles of <10° for substrates prior to incubation as shown inTable 1), suggesting that some oil may have been lost from the SLIPSduring those experiments. Additional control experiments demonstratedthis decrease in droplet sliding angles to result from incubation at thehigh densities of bacteria required for these pyocyanin assays(incubation under static conditions at lower densities of bacteria orshaking in the absence of bacteria did not affect droplet slidingangles). It has been reported in past studies that exposure to highshear forces (induced by flow, etc.) can promote the leaching of theinfused liquid phases of SLIPS in ways that can impact their slipperyproperties (see Howell, C., Vu, T. L., Johnson, C. P., Hou, X., Ahanotu,O., Alvarenga, J., Leslie, D. C., Uzun, O., Waterhouse, A., Kim, P.,Super, M., Aizenberg, M., Ingber, D. E., and Aizenberg, J. (2015)Stability of Surface-Immobilized Lubricant Interfaces under Flow, Chem.Mater. 27, 1792-1800; Howell, C., Vu, T. L., Lin, J. J., Kolle, S.,Juthani, N., Watson, E., Weaver, J. C., Alvarenga, J., and Aizenberg, J.(2014) Self-Replenishing Vascularized Fouling-Release Surfaces, ACSAppl. Mater. Inter. 6, 13299-13307.) It is also possible in this currentcontext that the presence of amphiphilic molecules produced by thebacterial cultures used here (i.e., lipids, lipid assemblies, etc.)could help promote the extraction of small amounts of oil over thecourse of these experiments. In support of this proposition, we notethat sliding angles could be restored to values of 10° or less by addingsmall amounts of silicone oil to the surface of the coatings (e.g., bypipette). Past reports also demonstrate that the gradual loss of infusedoil can be addressed in other practical ways, including through thedesign of porous substrates with oil reserves that can continuallyreplenish lost oil (Howell, C., Vu, T. L., Lin, J. J., Kolle, S.,Juthani, N., Watson, E., Weaver, J. C., Alvarenga, J., and Aizenberg, J.(2014) Self-Replenishing Vascularized Fouling-Release Surfaces, ACSAppl. Mater. Inter. 6, 13299-13307). Despite the increase in waterdroplet sliding angles observed in these experiments, the results ofadditional experiments described below demonstrate that thesecompound-loaded SLIPS-coated substrates remain anti-fouling to bacteriaunder similar culture conditions and can prevent the formation ofbacterial biofilms.

DMABI-Loaded SLIPS reduce biofilm formation on surrounding uncoatedsurfaces. As described above, the native silicone oil-infused SLIPS usedin this study have been demonstrated to resist the formation of P.aeruginosa biofilms under static culture conditions (e.g., in theabsence of added agents). To demonstrate the potential ofcontrolled-release SLIPS to also prevent biofilm formation insurrounding environments—and, thus, also confer measures ofanti-biofouling protection to nearby surfaces that are notSLIPS-coated—studies using SLIPS loaded with the anti-biofilm agentDMABI were also performed.

For these studies, P. aeruginosa was cultured in 12-well microtiterplates containing DMABI-loaded SLIPS substrates in the well bottoms(such that they were completely submerged in media). After 24 hours ofincubation, the amount of surface-attached biofilm on both the SLIPSsubstrates and the surrounding (uncoated) areas of the well bottoms wascharacterized by fluorescence microscopy and by staining with crystalviolet (CV). The SLIPS substrates were highly resistant to biofilmattachment, as expected from past studies (Manna et al., “Slipperyliquid-infused porous surfaces that prevent microbial surface foulingand kill non-adherent pathogens in surrounding media: A controlledrelease approach,” Advanced Functional Materials, 2016,26(21):3599-3611). No biofilm was observed over the entire centralregion of the coated substrates by fluorescence microscopy, and littleto no CV staining on the SLIPS surface (FIG. 19 (A-B)).

When the amount of CV on the SLIPS was quantified by UV/visspectrophotometry, more CV was observed than expected by visualinspection (corresponding to an approximately 50% reduction in staining;FIG. 19 (D)). It was determined, using fluorescence microscopy, thatthis residual staining was a result of the presence of biofilm near theuncoated edges of the SLIPS-coated substrates (FIG. 19 (F)), and not aresult of biofilm on the SLIPS surfaces themselves. These uncoated edgesare a result of the manner in which the SLIPS-coated substrates wereprepared for these proof-of-concept studies (e.g., by the fracture oflarger ‘parent’ SLIPS-coated surfaces into smaller ‘daughter’ chips; seethe above experimental sections for details) and are not an inherentlimitation of the SLIPS surfaces themselves.

As expected, native (unloaded) SLIPS had no significant influence on theformation of P. aeruginosa biofilms in regions of the surrounding(uncoated) well bottoms in these studies (FIG. 19 (C,E)). DMABI-loadedSLIPS, however, prevented biofilm attachment on the SLIPS-coated surfaceby CV staining (FIG. 19 (B,D)) and inhibited biofilm formation on thesurrounding uncoated well bottoms by approximately 50% (FIG. 19 (C,E))—aresult that is attributed to the gradual release of imbedded DMABI intosurrounding media (consistent with results shown in FIG. 17). It islikely that the levels of inhibition observed on uncoated well bottomsin these experiments (and the observation of persistent biofilm on theuncoated edges of the SLIPS-coated substrates, as noted above) could beimproved further by increasing the loading of DMABI or tuning the rateat which it is released through modifications to the structure of thepolymer matrix of the properties of the infused oil (the extendedrelease profile shown in FIG. 17 suggests that only small amounts ofDMABI were likely to have been released from these silicone oil-infusedSLIPS over the course of these short-term experiments). Efforts tooptimize and completely eradicate biofilm formation in the model systemused here were not pursued as part of these proof-of-concept studies,but would be straightforward to implement during the development orevaluation of these materials in application-specific contexts orenvironments.

Summary and Conclusions. Materials and surface coatings that areresistant to bacterial colonization and that can simultaneously inhibitbacterial virulence phenotypes on and around their surfaces would beuseful in a range of biomedical, environmental, and industrial contexts.Materials that can accomplish both of these important tasks withoutimpacting bacterial growth would be particularly valuable, as they wouldalso have the potential to avoid serious problems associated with thedevelopment of evolved resistance that currently plague traditionalbactericidal approaches. The work described here provides such materialsby developing new slippery and anti-fouling oil-infused surfaces thatcan be used as a robust platform for the controlled release or deliveryof small-molecule QSIs and biofilm inhibitors. These results demonstratethat this novel approach can significantly (i) reduce production of avirulence factor by planktonic bacteria in the vicinity of the surfaceand (ii) reduce the biofilm burden on the surface of the material itselfand on surrounding non-SLIPS-coated surfaces. These results alsodemonstrate that this controlled-release SLIPs approach can be used toload and release of combinations or ‘cocktails’ of these agents that maybe more effective than any single antibiotic or QSI alone.

The methods used to fabricate these slippery coatings can be used tocoat topologically complex substrates, including tubing, filters, orimplants, and should thus allow for protection against surface foulingand facilitate the local, controlled delivery of anti-virulence agentsdirectly to sites endemic to bacterial colonization in medical devicesand/or industrial equipment. The modular nature of these SLIPS alsoprovides opportunities to tune the slippery and controlled releaseproperties of these coatings though changes in the structure of thepolymer matrix, the physicochemical properties of the infused oil phase,and the solubilities and structures of the small-molecule agents thatare loaded. The approaches and new strategies described here can thusform the basis of a general and multi-functional materials platform thatare useful for combating bacterial biofouling and virulence vianon-biocidal pathways in a range of important fundamental and appliedcontexts.

Example 5—Design of Controlled Release SLIPS Loaded withTertamethylrhodamine

Experiments were conducted comparing the release rates oftertamethyl-rhodamine (TMR) from SLIPS prepared by infusing a porouspolymer matrix with a saturated solution of TMR in silicone oil, andSLIPS prepared by first loading TMR directly into a polymer matrix andthen infusing with the silicone oil.

A saturated solution of tertamethylrhodamine (TMR) in silicone oil wasfirst prepared by adding 0.5 mg of TMR to 20 mL of silicone oil,maintaining it on a shaker plate for 24 hours, and then removingundissolved TMR by centrifugation at 5000 rmp for 2 min. 3 μL of thisTMR/silicone oil solution was then spread on and infused into driednanoporous PEI/PVDMA multilayers. The release of TMR was characterizedby incubating these TMR-loaded SLIPS in PBS. A release curve arisingfrom samples prepared in this way is provided in FIG. 20 (A) and showsthat release occurred rapidly.

Next, SLIPS were prepared by loading the TMR onto the polymer matrixbefore addition of the oil. Briefly, 10 μL of a solution of TMR inacetone (0.02 mg/mL) was placed on dried nanoporous PEI/PVDMAmultilayers and allowed to dry as described in previous examples. 3 μLof silicone oil was then spread on and infused into the polymer matrix.The release of TMR was characterized by incubating these TMR-loadedSLIPS in PBS. A release curve arising from samples prepared in this wayis provided in FIG. 20 (B) and shows (i) that release occurred moreslowly than the example in which TMR was loaded in the oil phase only(FIG. 20 (A)), and (ii) the loading was higher using this method,because this approach circumvents limitations related to the solubilityof TMR in silicone oil and allows greater amounts of TMR to be loadedprior to oil infusion.

Having now fully described the present invention in some detail by wayof illustration and examples for purposes of clarity of understanding,it will be obvious to one of ordinary skill in the art that the same canbe performed by modifying or changing the invention within a wide andequivalent range of conditions, formulations and other parameterswithout affecting the scope of the invention or any specific embodimentthereof, and that such modifications or changes are intended to beencompassed within the scope of the appended claims.

One of ordinary skill in the art will appreciate that startingmaterials, reagents, purification methods, materials, substrates, deviceelements, analytical methods, assay methods, mixtures and combinationsof components other than those specifically exemplified can be employedin the practice of the invention without resort to undueexperimentation. All art-known functional equivalents, of any suchmaterials and methods are intended to be included in this invention. Theterms and expressions which have been employed are used as terms ofdescription and not of limitation, and there is no intention that theuse of such terms and expressions exclude any equivalents of thefeatures shown and described or portions thereof, but it is recognizedthat various modifications are possible within the scope of theinvention claimed. Thus, it should be understood that although thepresent invention has been specifically disclosed by preferredembodiments and optional features, modification and variation of theconcepts herein disclosed may be resorted to by those skilled in theart, and that such modifications and variations are considered to bewithin the scope of this invention as defined by the appended claims.

As used herein, “comprising” is synonymous with “including,”“containing,” or “characterized by,” and is inclusive or open-ended anddoes not exclude additional, unrecited elements or method steps. As usedherein, “consisting of” excludes any element, step, or ingredient notspecified in the claim element. As used herein, “consisting essentiallyof” does not exclude materials or steps that do not materially affectthe basic and novel characteristics of the claim. In each instanceherein any of the terms “comprising”, “consisting essentially of” and“consisting of” may be replaced with either of the other two terms.

When a group of materials, compositions, components or compounds isdisclosed herein, it is understood that all individual members of thosegroups and all subgroups thereof are disclosed separately. When aMarkush group or other grouping is used herein, all individual membersof the group and all combinations and subcombinations possible of thegroup are intended to be individually included in the disclosure. Everyformulation or combination of components described or exemplified hereincan be used to practice the invention, unless otherwise stated. Whenevera range is given in the specification, for example, a temperature range,a time range, or a composition range, all intermediate ranges andsubranges, as well as all individual values included in the ranges givenare intended to be included in the disclosure. In the disclosure and theclaims, “and/or” means additionally or alternatively. Moreover, any useof a term in the singular also encompasses plural forms.

All references cited herein are hereby incorporated by reference intheir entirety to the extent that there is no inconsistency with thedisclosure of this specification. Some references provided herein areincorporated by reference to provide details concerning sources ofstarting materials, additional starting materials, additional reagents,additional methods of synthesis, additional methods of analysis,additional biological materials, and additional uses of the invention.All headings used herein are for convenience only. All patents andpublications mentioned in the specification are indicative of the levelsof skill of those skilled in the art to which the invention pertains,and are herein incorporated by reference to the same extent as if eachindividual publication, patent or patent application was specificallyand individually indicated to be incorporated by reference. Referencescited herein are incorporated by reference herein in their entirety toindicate the state of the art as of their publication or filing date andit is intended that this information can be employed herein, if needed,to exclude specific embodiments that are in the prior art. For example,when composition of matter are claimed, it should be understood thatcompounds known and available in the art prior to Applicant's invention,including compounds for which an enabling disclosure is provided in thereferences cited herein, are not intended to be included in thecomposition of matter claims herein.

1.-20. (canceled)
 21. An anti-biofouling surface able to controllablyrelease one or more molecules, wherein said anti-biofouling surfacecomprises: a) a porous matrix having nanoscale or microscale porosity;b) an oil covering at least a portion of the porous matrix, wherein saidoil at least partially fills the pores of the porous matrix, whereinsaid oil is selected from the group consisting of a silicone oil, athermotropic liquid crystal, and combinations thereof; and c) one ormore releasable molecules, wherein the one or more releasable moleculesare located on the surface or in said porous matrix, within said oil, orboth, wherein the anti-biofouling surface is able to controllablyrelease the one or more releasable molecules when the anti-biofoulingsurface is immersed into an aqueous media, and wherein the one or morereleasable molecules are able to reduce, inhibit, or modulate thebehaviors of non-adherent pathogens in the aqueous media.
 22. Theanti-biofouling surface of claim 21 wherein the oil comprises athermotropic liquid crystal.
 23. The anti-biofouling surface of claim21, wherein the non-adherent pathogens are selected from the groupconsisting of Candida albicans, Pseudomonas aeruginosa, Escherichiacoli, Staphylococcus aureus, and combinations thereof.
 24. Theanti-biofouling surface of claim 21 wherein the one or more releasablemolecules comprise a natural or synthetic antibiotic agent, a natural orsynthetic antifungal agent, an agent that modulates bacterial or fungalquorum sensing, an agent that attenuates virulence, or a combinationthereof.
 25. The anti-biofouling surface of claim 24 wherein the one ormore releasable molecules is selected from the group consisting oftriclosan, acyl L-homoserine lactone (AHL) derivatives,aminobenzimidazole (ABI) derivatives, and combinations thereof.
 26. Theanti-biofouling surface of claim 21 wherein the one or more releasablemolecules are selected from the group consisting of:

or combinations thereof.
 27. The anti-biofouling surface of claim 21wherein the porous matrix comprises a multilayer film having one or morebilayers, wherein each bilayer comprises a first polymer layer incontact with a second polymer layer, where said multilayer film hasnanoscale or microscale porosity.
 28. The anti-biofouling surface ofclaim 27 wherein the first polymer layer comprises a functionalizedazlactone having the formula:

wherein x is 0 or the integers 1 or 2; and each R¹ is independentlyselected from the group consisting of: hydrogen, alkyl groups, alkenylgroups, alkynyl groups, carbocyclic groups, heterocyclic groups, arylgroups, heteroaryl groups, alkoxy groups, aldehyde groups, ether groups,and ester groups, any of which may be substituted or unsubstituted. 29.The anti-biofouling surface of claim 27 comprising one or more PVDMA/PEIbilayers, which are further functionalized with n-decylamine and whereinthe one or more bilayers are infused with a silicone oil or ananisotropic thermotropic liquid crystal.
 30. A container or hollow tubehaving an inner surface comprising the anti-biofouling surface of claim21.
 31. A catheter tube having an inner surface comprising theanti-biofouling surface of claim
 21. 32. A method for fabricating ananoporous or microporous anti-biofouling surface able to reduce,inhibit, or modulate the behaviors of non-adherent pathogens insurrounding aqueous media, said method comprising the steps of: a)forming a porous matrix on a substrate, wherein said porous matrix hasnanoscale or microscale porosity; b) exposing the porous matrix to anoil, wherein said oil coats at least a portion of the porous matrix andsaid oil at least partially fills the pores of at least a portion ofsaid porous matrix, wherein the oil is selected from the groupconsisting of a silicone oil, a thermotropic liquid crystal, andcombinations thereof; and c) loading one or more releasable moleculesonto said porous matrix or into said oil, wherein the one or morereleasable molecules comprise a natural or synthetic antibiotic agent, anatural or synthetic antifungal agent, an agent that modulates bacterialor fungal quorum sensing, an agent that attenuates virulence, or acombination thereof, and wherein the anti-biofouling surface is able tocontrollably release the one or more releasable molecules when theanti-biofouling surface is immersed into the aqueous media.
 33. A methodfor reducing, inhibiting, or modulating the behaviors of non-adherentpathogens in an aqueous media surrounding a substrate comprising thesteps of: a) providing an anti-biofouling surface on the substrate, saidanti-biofouling surface comprising: i) a porous matrix having nanoscaleor microscale porosity; ii) an oil covering at least a portion of theporous matrix, wherein said oil at least partially fills the pores ofthe porous matrix, wherein the oil is selected from the group consistingof a silicone oil, a thermotropic liquid crystal, and combinationsthereof; and iii) one or more releasable molecules able to reduce,inhibit, or modulate the behaviors said non-adherent pathogens uponcontact with said non-adherent pathogens, wherein the one or morereleasable molecules are located on the surface of said porous matrix,within said oil, or both; b) controllably releasing the one or morereleasable molecules from said anti-biofouling surface into said aqueousmedia, wherein the one or more releasable molecules contact thenon-adherent pathogens thereby reducing the number of non-adherentpathogens, inhibiting the growth or colonization of the pathogens, ormodulating the behaviors of the non-adherent pathogens.
 34. The methodof claim 33 wherein the non-adherent pathogens are selected from thegroup consisting of Candida albicans, Pseudomonas aeruginosa,Escherichia coli, Staphylococcus aureus, and combinations thereof. 35.The method of claim 33 wherein the one or more releasable moleculescompounds is a natural or synthetic antibiotic agent, natural orsynthetic antifungal agent, quorum sensing modulator, or a combinationthereof.
 36. The method of claim 33 wherein the one or more releasablemolecules is selected from the group consisting of of triclosan, acylL-homoserine lactone (AHL) derivatives, aminobenzimidazole (ABI)derivatives, and combinations thereof.
 37. The method of claim 33wherein the one or more releasable molecules is selected from the groupconsisting of:

or combinations thereof.
 38. The method of claim 33 wherein the porousmatrix comprises a multilayer film having one or more bilayers, whereineach bilayer comprises a first polymer layer in contact with a secondpolymer layer, where said multilayer film has nanoscale or microscaleporosity.
 39. The method of claim 33 wherein the anti-biofouling surfacecomprises one or more PVDMA/PEI bilayers, which are furtherfunctionalized with n-decylamine and wherein the one or more bilayersare infused with a silicone oil or an anisotropic thermotropic liquidcrystal.
 40. The method of claim 33 further comprising the step ofloading an additional one or more releasable molecules within the oilwhen levels of releasable molecules drop below a desired level.